Abstract
Cytogenetics, with its fundamental role in the field of genetic investigation, continues to be an indispensable tool for studying phylogenetics, given that currently molecular evolutionary analyses are more commonly utilized. Chromosomal evolution indicated that genomic evolution occurs at the level of chromosomal segments, namely, the genomic blocks in the size of Mb‐level. The recombination of homologous blocks, through the mechanisms of insertion, translocation, inversion, and breakage, has been proven to be a major mechanism of speciation and subspecies differentiation. Meanwhile, molecular cytogenetics (fluorescence in situ hybridization‐based methodologies) had been already widely applied in studying plant genetics since polyploidy is common in plant evolution and speciation. It is now recognized that comparative cytogenetic studies can be used to explore the plausible phylogenetic relationships of the extant mammalian species by reconstructing the ancestral karyotypes of certain lineages. Therefore, cytogenetics remains a feasible tool in the study of comparative genomics, even in this next generation sequencing (NGS) prevalent era.
Keywords
- cytogenetics
- comparative cytogenetics
- fluorescence in situ hybridization
- genomic insitu hybridization
- zoo‐CGH
1. Introduction: chromosomal evolution of mammals
According to fossil records, the radiation evolution of mammals diverged after the K‐T boundary (approximately 65 Mya, between the Cretaceous and Tertiary periods, at which most of the dinosaurs were extinct). There are three hypotheses that try to explain such findings: (1) Explosive hypothesis: It is supported by most paleobiologists and states that the genesis and diversification of many phyletic groups (“Orders”) diverged after the Cretaceous-Tertiary (K-T) boundary; (2) Long Fuse hypothesis: It supports the view that Order diversification occurred after the K‐T boundary but that genesis occurred in the Cretaceous period, i.e., before the K‐T boundary; and (3) Short Fuse hypothesis: It considers the genesis and diversification of Orders to have diverged before the K‐T boundary (Figure 1) [1].
Molecular data indicate that mammalian diversification began in the Cretaceous period, which supports the (2) Long Fuse and (3) Short Fuse hypotheses. However, these data have limitations, including the availability of a single temporal calibration point and the variable evolution rate of different phyletic groups. Due to the lack of representativeness of the samples, this inadequate taxon sampling restricts the use on some, but not all, placental mammals, and it makes the negative correlation between evolution rate and body size difficult to explain. William Murphy and Stephen O’Brien’s team made a successful attempt at answering these questions with zoo‐fluorescence
Figure 2 presents the phylogenetic tree of placental mammals derived from 16,379 nucleotide sequences (including 19 nuclear genes and 3 mitochondrial genes published by the study team), where opossum is considered an outgroup using the maximal likelihood method, and placental mammals are considered to appear at 105 Mya. When the K‐T boundary is labeled with red dashes, we find that “Order” genesis and diversification are events that occur before the boundary.
By comparing the chromosomal break point of multiple species, including the chromosomal rearrangement of loci discovered via comparative genomics and some genetic sequences from fully sequenced species, we can clearly find that (1) Approximately 20% of chromosomal break points are repeatedly involved in the evolutionary process of mammals. (2) These repeatedly involved break points are primarily located at the centromere and telomere. (3) The number of genes within and near the break point blocks that are involved in chromosomal evolution is higher than the mean of the overall genome. (4) The unique break points unique in Primates are located at repeated segment regions and the ends are surrounded by reversed sequences. Figure 3 refers to the rate of chromosomal breaking using the chromosomal break points involved in the evolution of mammals.
The result shows that the chromosomal rearrangement rate before the K‐T boundary is 0.11–0.43/My, and this rate is doubled to quadrupled for Primates and increased fivefold for Rodentia [2, 3].
2. How to apply molecular genomics in the study of evolution and parental relationships
2.1. Zoo‐FISH
Comparative mapping: It is a method for comparing the location of homologous genes of different species to explore the evolution of genomes; zoo‐FISH is an extension of such technology. This method assesses the overall chromosomal similarity among all mammalian orders and becomes a powerful tool to study genomic evolution. The possible mechanism and factors related to mammalian genomic evolution can be understood with Metatheria and Eutheria studies.
When conducting zoo‐FISH, partial or whole chromosomes are obtained through the sorting of fluorescence‐labeled cells or microscopic extraction. DNA extracted from this specific chromosomal block is subject to degenerated oligonucleotide primed‐PCR (DOP‐PCR), then labeled with fluorescence to produce probes, and hybridized with the chromosome of the species of interest. Due to the resolution of zoo‐FISH, which is approximately 10 Mbp (megabase pairs), this method may underestimate the real rearrangement events on the chromosome. However, zoo‐FISH has revealed some interesting facts: many chromosome blocks of different species are rather conservative, and the similar chromosome blocks from a common ancestor are called synteny blocks. For example, one somatic chromosome of the gray‐headed flying fox (
One of the most important applications of zoo‐FISH is to study the speed of chromosomal rearrangement when studying genomic evolution [5]. Using the phylogenetic tree that is based on fossil evidence, we can understand the rate of movement and rearrangement of synteny blocks in the chromosomes of two species. When there are difficulties in bi‐directional zoo‐FISH, monodirectional zoo‐FISH can provide with key information or a new understanding. By comparing the chromosomal synteny blocks of indicator mammals and Aves, the occurrence rate of chromosomal rearrangement was found to be fixed at approximately 1–10/Mya [6]. The chromosomal rearrangement rate is shown in Figure 4, and the rate may differ with lineage genesis and at different evolutionary stages.
Three important stages of chromosomal rearrangement are found (Figure 4): The first stage (1–3 Mya) < 0.2/My, the second stage increased to 1.1/My, and in the third stage, the rearrangement rate greatly varied in nonrodents. For example, humans,
The other application of zoo‐FISH is to reconstruct primitive karyotyping. Figure 6 shows the estimates of ancestral placental mammal (2
It is worth noting that the study shows that the chromosomal karyotype of primitive placental mammals is 2
The karyotype of Hoffmann’s two‐toed sloth: The blocks that are syntenic to HSA are labeled on the left of each chromosome. For example, Chromosome 1 is syntenic to HSA1, but it is not syntenic to other HSA chromosomes, while Chromosome 6 contains synteny blocks that are similar to those found in HSA3 and HSA21 [7]. These karyotypes are presented in Figure 7.
2.2. How is chromosomal recombination fixed in evolution?
Theoretically, chromosomal rearrangement may lead to meiotic errors and reduced fertility. It is fundamentally a harmful genetic variation, and most rearrangements are difficult to pass on in a population. However, (1) genetic drift, (2) Muller’s ratchet mechanism or (3) hitchhiker make it possible to keep some chromosomal recombination (beneficial mutations may be eliminated due to the selection of other loci, whereas harmful mutations may be preserved due to the selection of other beneficial loci).
2.3. The importance of studying “the weird mammals”
The genome of most mammals contains approximately 3 billion nucleotides (3 × 109 bp), but the number of chromosome varies greatly. For placental mammals, Indian muntjac possesses as few as 2
Infraclasses Eutheria (placental mammals) and Metatheria (opossum) diverged at approximately 130 Mya, and their subclasses, Theria and Prototheria (i.e., monotreme), diverged at approximately 170 Mya. Fossil studies show that the radiation evolution of placental mammals (20 orders, including more than 4600 species) occurred in the Cretaceous period (approximately 60–80 Mya). By comparing the differences in the genomes of various animal populations, especially those that play specific roles in evolutionary history (Jennifer Graves, an Australian scholar, called them “the weird mammals”), such as monotreme, opossum and fast‐evolving rodents, we can learn more about the evolution progress of mammals.
3. The innovative application of zoological comparative genomic hybridization (CGH) in phylogenetics
Placental mammals include four major lineages: (1) Afrotheria, which includes the orders Sirenia, Hyracoidea, Proboscidea, Tubulidentata, Macroscelidea and Afrosoricida; (2) Laurasiatheria, which includes orders Eulipotyphla, Carnivora, Pholidota, Perissodactyla, Cetartiodactyla and Chiroptera; (3) Euarchontoglires, which includes Rodentia, Lagomorpha, Primates, Scandentia and Dermoptera; and (4) Xenarthra [8]. Currently, there are disputes and uncertainties in the phylogenetic relationships and the true origins of each order in these four lineages. We attempt to define the phylogenetic relationship of the orders Pholidota, Carnivora and Xenarthra using genomic
In this “DNA‐DNA hybridization,” the DNA of two species was cut into small chunks of 600–800 bp before mixing. Unfortunately, this technology was unable to prevent errors that were caused by the existence of paralogous sequences instead of orthologous sequences. The result was used for trending, similar to zoo‐GISH, but it was not designed for accuracy. On the other hand, analyses that are focused on one or more genes that are present in the evolutionary history of only a few loci, lack a bridge to connect them. We are looking for a tool that is capable of not only whole genome and individual gene trending, but also larger block trending for genomes, and even positioning. Therefore, the author chose to apply a mature technology from the study of human neoplasms called “metaphase comparative genomic hybridization (CGH)” to the study of phylogenetic history.
4. The history and prior applications of CGH
In 1992, Dan Pinkel’s lab at UC San Francisco published an innovative technology named CGH [10]. In this method, tumor and normal cellular DNA probes were labeled with red and green fluorescence, respectively. They were then hybridized with normal cells in metaphase and competed with each other in incorporating with normal chromosomes. Yellow is observed when red and green fluorescence are mixed in equal amounts. A block with more tumor cell genome than the normal reference, i.e., with duplication, turns green, whereas deletion causes it to turn red. This innovative genome‐wide technology not only allows positioning, but shows increase or decrease, making it a powerful tool in searching for tumor suppressing genes (which make the amount of tumor cells lower than those of normal reference) or oncogenes (which make the amount of tumor cells higher than those of normal reference), with a resolution of 5–10 Mbp. However, this technology is difficult to operate and requires specific photographic tools and image processing software to calculate the ratio of red and green fluorescence. Recently, gene chips have replaced this technology. Gene chips, formally known as array CGH (the original CGH was renamed as metaphase CGH), have designated probes that are fixed onto a chip [11]. The array CGH probes are derived from the known sequences of target organisms. Array CGH does not involve chromosomal preparation or microscope interpretation. Conversely, metaphase CGH is genome‐wide and has chromosome‐level resolution, and it is a useful tool when the full genome sequence is unknown. This technology can be applied in more than tumor research; it is also valuable for studying human genetic diseases that are related to repeated or deleted blocks, especially those that are caused by copy number variation [12]. The captured images and the last interpretation are presented in Figure 9, where (A) fluorescein (FITC) is used to provide green light; (B) rhodamine for red light; and (C) merged CGH results from one normal sample.
The fluorescence of the green‐red ratio was analyzed with software.
We also applied this technology to report a rare case of missing human 13q31 without clinical symptoms [13]. In Figure 10, we can see that the human 13q31 block presents more red fluorescence in the block indicated by a straight red line (considered an increase when the green‐red ratio is more than 1.2 and a decrease when the ratio is less than 0.8). The label
Based on the experience of metaphase CGH in human medicine, the author considered the feasibility of applying this technology in interspecies exploration to characterize the evolutionary relationships among extant eutherian mammalian taxonomic groups (orders/supraordinal clades). That is, to determine the sequence/genomic similarity of unknown‐sequence species A and B with respect to species C, the DNA of species A and B would be labeled with molecules emitting different fluorescence dyes. The ratio of the labeled fluorescence intensities in each chromosome of species C should then reflect regions of sequence similarity to species A versus B. This is a brand‐new application and the author named it “zoo‐CGH” (Figure 11).
5. Applying CGH in exploring the relationship between Pholidota , Carnivora , and Xenarthra
Myrmecophagy is a feeding behavior characterized by mainly or exclusively eating ants, termites, or both. This feeding specialization occurs in few eutherian mammals. Myrmecophagous species of Eutheria are in the orders Pholidota (e.g., pangolins,
5.1. Method and procedures
5.1.1. Determine nuclear genome size
The genome size of the two‐toed sloth and domestic dog were determined to ensure that approximately equal numbers of nuclei (i.e., copy number of whole genomes in each species) are used in zoo‐CGH analyses. The genome sizes were obtained after flow cytometry analysis of propidium iodide (IP)‐stained nuclei from the target organisms.
5.1.2. Extract DNA from the two‐toed sloth and domestic dog
Genomic DNA was isolated from leukocytes with a commercial kit (Gentra Puregene DNA Purification Kit, Qiagen, Hilden, German), used in accordance with the manufacturer’s instructions.
5.1.3. Prepare the mitotic metaphase slides of Taiwanese pangolin
Fibroblast cell lines were established from lung tissues derived from Taiwanese pangolin, and metaphase cells were harvested following a 2‐hour incubation with colcemid (at a concentration of 0.1 μg/ml).
5.1.4. Produce two‐toed sloth and domestic dog DNA probes
The two‐toed sloth and domestic dog DNA were labeled with biotin and digoxigenin (DIG) by nick translation, respectively.
5.1.5. Prepare pangolin C0t‐1 DNA
5.1.6. Perform zoo‐CGH
Male Taiwanese pangolin chromosome spreads were prepared on a slide and denatured at 73°C for 5 minutes in 70% formamide and 2 × SSC, pH 7.0, followed by dehydration in a graded ethanol series. Next, equal genome copy numbers of biotin‐labeled two‐toed sloth DNA and DIG‐labeled domestic dog DNA were coprecipitated with a 50‐fold excess of Taiwanese pangolin
5.1.7. Analyze image
By comparing the fluorescence ratio on the longitudinal axis of pangolin metaphase chromosome, we estimated differences in the inter‐species gene copy number and DNA sequence similarity. The means of the F/R ratios obtained from the heterologous hybridization, which represents DNA from different species labeled with different fluorophores that are competitively bound to probes obtained from a third species, were calculated for each pangolin autosome. Pangolin chromosomal segments with F/R ratios of < 0.8 (red fluorescence is more intense) and > 1.2 (green fluorescence is more intense) were considered to have significantly different hybridization strengths. When the F/R ratios were between 0.8 and 1.2 (showing yellow fluorescence), the DNA sequence difference or copy number of each pair was roughly equivalent. Means of the ratios were also calculated using a dye‐swap design.
5.2. Result
In Figure 12, we can see red, green or yellow blocks on different parts of the chromosome. The overall homology between the pangolin and dog genomes was higher than that between the pangolin and sloth genomes. Analysis of pangolin chromosomes 14 and 15, which were the largest and most easily identifiable, showed that red fluorescence is dominant in euchromatin, i.e., more similar to the domestic dog (Figure 12E). When dye swapping was conducted, i.e., green fluorescence for the domestic dog and red fluorescence for the two‐toed sloth, consistent results were obtained (Figure 12F).
Figure 12 shows zoo‐CGH for the domestic dog, two‐toed sloth, and Taiwanese pangolin. In panel (A) genomic DNA from dog (labeled with DIG conjugated to the red fluorophore, rhodamine) and sloth (labeled with biotin conjugated to the green fluorophore, fluorescein) were mixed in equal quantities and competitively hybridized to metaphase spreads from the pangolin lymphocytes. In panel (B) individual chromosome analysis of the fluorescent ratio in (A) was presented where blue lines denote the ratio of F/R signal at each position of the pangolin chromosomes. Numbers in brackets represent the number of chromosomes analyzed. When the vertical bar between each chromosome and its ideogram appears red or green, the F/R ratio was <0.8 or >1.2, respectively. Overall, all chromosomes (except Y) appeared red. Panels (C) and (D) represent dye swap of (A) and (B), respectively. All chromosomes (except Y) appeared green.
From the results above, we found that all somatic chromosomes of
6. Discussion
In early times, comparative genomics study between closely related species can only be done by comparing the karyotypes of the species and the techniques used are primitive, including Giemsa stain only, the G‐banding techniques, and thus only the diploid number (2N), the functional number (FN, indicating the numbers of the chromosomal arms), as well as the classification of the chromosomes into metacentric, submetacentric, acrocentric, and telocentric according to the arm ratios can be provided. In addition, the special stains, such as the C‐banding and Ag-nucleolus organizer region (NOR) staining, can be used to elucidate the constitutive heterochromatin (by C‐banding), and the sites of secondary constriction and the active‐transcribing ribosomal DNA genes (by Ag‐NOR staining), can help to find the more trivial differences between species which may carry evolutionary significance [18, 19]. However, the advent of fluorescence
7. Conclusion
Despite molecular evolution being made nowadays, by studying the homologous DNA sequences and using different evolutionary analytical models to reconstruct the phylogeny, which is the mainstream of comparative genomics [1–4], especially when sequencing the whole genome of each species has become more feasible through the powerful next generation sequencing (NGS) technology [21], cytogenetics remains an indispensible tool in studying the karyotypic evolution, which is one of the major mechanisms and thus is equally important as the molecular evolution to the processes involved in the speciation and subspecies differentiation. Conventional karyotyping, special stains to delineate the locations of heterochromatin, sites of active‐transcribing ribosomal DNA genes, as well as molecular cytogenetics (namely, the fluorescence
References
- 1.
Springer MS, Murphy WJ, Eizirik E, O’Brien SJ. Placental mammal diversification and the Cretaceous‐Tertiary boundary. Proceedings of the National Academy of Sciences of the United States of America. 2003;100:1056–1061. DOI: 10.1073/pnas.0334222100 - 2.
Murphy WJ, Eizirik E, Johnson WE, Zhang YP, Ryder OA, O’Brien SJ. Molecular phylogenetics and the origins of the placental mammals. Nature. 2001;409:614–618. DOI: 10.1038/35054550 - 3.
Murphy WJ, Larkin DM, Wind AE, Bourque G, Tesler G, Auvil L, Beever JE, Chowdhary Bp, Galibert F, Gatzke L, Hitte C, Meyers SN, Milan D, Ostrnder EA, Pape G, Parker HG, Raudsepp T, Rogatcheva MB, Schook LB, Skow LC, Welge M, Womack JE, O’Brien SJ, Pevzner PA, Lewin HA. Dynamics of mammalian chromosome evolution inferred from multispecies comparative maps. Science. 2005;309:613–617. DOI: 10.1126/science.1111387 - 4.
Murphy WJ, Stanyon R, O’Brien SJ. Evolution of mammalian genome organization inferred from comparative gene mapping. Genome Biology. 2001b;2(6):reviews0005.1‐reviews0005.8. DOI: 10.1186/gb‐2001‐2‐6‐reviews0005 - 5.
Ferguson‐Smith MA, Yang F, RensW, O’Brien PC. The impact of chromosome sorting and painting on the comparative analysis of primate genomes. Cytogenetic and Genome Research. 2005;108(1–3):112–121. DOI: 10.1159/000080809 - 6.
Burt DW, Bruley C, Dunn IC, Jones CT, Ramage A, Law AS, Morrice DR, Paton IR, Smith J, Windsor D, Sazanov A, Fries R, Waddington D. The dynamics of chromosome evolution in birds and mammals. Nature. 1999;402:411–413. DOI: 10.1038/46555 - 7.
Svartman M, Stone G, Stanyon R. The ancestral eutherian karyotype is present in Xenarthra. PLoS Genetics. 2006;2(7):e109. DOI: 10.1371/journal.pgen.0020109 - 8.
Amrin‐Madsen H, Koepfli KP, Wayne RK, Springer MS. A new phylogenetic marker, Apolipoprotein B, provides compelling evidence for eutherian relationships. Molecular Phylogenetics and Evolution. 2003;28(2):225–240. DOI: 10.1016/S1055‐7903(03)00118‐0 - 9.
Sibley CG, Ahlquist JE. The phylogeny of the hominoid primates, as indicated by DNA‐DNA hybridization. Journal of Molecular Evolution. 1984;20(1):2–15. DOI: 10.1007/BF02101980 - 10.
Kallioniemi A, Kallioniemi OP, Sudar D, Rutovitz D, Gray JW, Waldman F, Pinkel D. Comparative genomic hybridization for molecular cytogenetic analysis of solid tumors. Science. 1992;258(5083):818–821. DOI: 10.1126/science.1359641 - 11.
Kallioniemi A. CGH microarrays and cancer. Current Opinion in Biotechnology. 2008;19(1):36–40. DOI: 10.1016/j.copbio.2007.11.004 - 12.
Lee C, Iafrate AJ, Brothman AR. Copy number variations and clinical cytogenetic diagnosis of constitutional disorders. Nature Genetics. 2007;39(7 Suppl):S48‐S54. DOI: 10.1038/ng2092 - 13.
Ke YY, Lee DJ, Ma GC, Lee MH, Wang BT, Chen M. Interstitial deletion 13q31 associated with normal phenotype: cytogenetic study of a family with concomitant segregation of reciprocal translocation and interstitial deletion. Journal of the Formosan Medical Association. 2007;106(7):582–588 DOI: 10.1016/S0929‐6646(07)60010‐2 - 14.
McNab BK. Physiological convergence amongst ant‐eating and termite‐eating mammals. Journal of Zoology. 1984;203:485–510. DOI: 10.1111/j.1469‐7998.1984.tb02345.x - 15.
Redford KH. Ant and termites as food: Patterns of mammalian myrmecophagy. In: Genoways HH, editor. Current Mammalogy. New York: Springer; 1987. pp. 349–399. DOI: 10.1007/978‐1‐4757‐9909‐5_9 - 16.
Yu HT, Ma GC, Lee DJ, Chin SC, Tsao HS, Wu SH, Shih SY, Chen M. Molecular delineation of the Y‐borne Sry gene in the Formosan pangolin ( Manis pentadactyla pentadactyla ) and its phylogenetic implications for Pholidota in extant mammals. Theriogenology. 2011;75(1):55–64. DOI: 10.1016/j.theriogenology.2010.07.010 - 17.
Yu HT, Ma GC, Lee DJ, Chin SC, Chen TL, Tsao HS, Lin WH, Wu SH, Lin CC, Chen M. Use of a cytogenetic whole‐genome comparison to resolve phylogenetic relationships among three species: Implications for mammalian systematics and conservation biology. Theriogenology. 2012;77(8):1615–1623. DOI: 10.1016/j.theriogenology.2011.12.006 - 18.
Wu SH, Chen M, Chin SC, Lee DJ, Wen PY, Chen LW, Wang BT, Yu HT. Cytogenetic analysis of the Formosan pangolin Manis pentadactyla pentadactyla (Mammalia: Pholidota). Zoological Studies. 2007;46:389–396 - 19.
Lin LK, Ma GC, Chen TH, Lin WH, Lee DJ, Wen PY, Wu SH, Chen M. Genomic analyses of the Formosan harvest mouse (Micromys minutus) and comparisons to the brown Norway rat (Rattus norvegicus) and the house mouse (Mus musculus). Zoology (Jena). 2013;116(5):307–315. DOI: 10. 1016/j.zool.2013.07.001 - 20.
Silva GS, Souza MM. Genomic in situ hybridization in plants. Genetics and Molecular Research 2013;12(3):2953–2965. DOI: 10.4238/2013.August.12.11 - 21.
Pennisi E. Sequencing all life captivates biologists. Science. 2017;355(6328):894–895. DOI: 10.1126/Science.355.6328.894