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This is reference to a review on C. elegans mosaic analysis by Yochem and Herman, in which the authors make a distinction between free chromosome fragments and extrachromosomal arrays.
For the former, they reference Herman 1984. For the latter, they reference Lackner et al 1994 and Miller et al 1996, but these are studies that use extrachromosomal arrays, not papers defining them as a technique, per se.
I am curious to know what the distinction is - I am under the impression that extrachromosomal arrays are fragments with many copies of the gene of interest + a marker.
I asked my professor, and the answer appears to be differences in both the generation and the final product.
Free chromosome fragments are created through irradiation/other damage of the germline in one animal. Through a series of crosses, it is possible to introduce individual fragments (containing a duplication of your gene of interest, as well as a marker) into a mutant background. The extrachromosomal fragment will be lost during mitosis at various points in development, and in a mosaic analysis, you can look for the loss of your marker (and therefore the loss of the wildtype allele), and see if yfg product is required in certain cell types for a wildtype phenotype.
Extrachromosomal arrays are created by cloning, and you typically have many copies of both the gene and marker. It's the same principle as free chromosomal fragments in terms of mosaic analysis, but it's very powerful, as you can easily clone individual genes (versus irradiating and hoping to discover yfg on a fragment) and markers (and now you can use non-endogenous markers like GFP).
Even though extrachromosomal arrays are comparatively easier to generate, the problem is that you end up with many copies of the gene, which leads to non-endogenous dosage levels. Even if this doesn't cause a completely different mutant phenotype in the cells, it is possible that enough of these proteins could segregate at mitosis into the daughter cells, such that even if the fragment is lost, you could still have your gene product around in the daughter cells (if your marker protein didn't also segregate like this, you might draw erroneous conclusions about the necessity of yfg product in those particular cells).
Identification of chromosome sequence motifs that mediate meiotic pairing and synapsis in C. elegans
Caenorhabditis elegans chromosomes contain specialized regions called pairing centres, which mediate homologous pairing and synapsis during meiosis. Four related proteins, ZIM-1, 2, 3 and HIM-8, associate with these sites and are required for their essential functions. Here we show that short sequence elements enriched in the corresponding chromosome regions selectively recruit these proteins in vivo. In vitro analysis using SELEX indicates that the binding specificity of each protein arises from a combination of two zinc fingers and an adjacent domain. Insertion of a cluster of recruiting motifs into a chromosome lacking its endogenous pairing centre is sufficient to restore homologous pairing, synapsis, crossover recombination and segregation. These findings help to illuminate how chromosome sites mediate essential aspects of meiotic chromosome dynamics.
SDC proteins function as a complex in vivo to repressher-1 and X chromosomes
How does SDC-2 discriminate between different targets and repress them to different degrees? To answer this question, we first identified proteins that function with SDC-2 at her-1. sdc-2 was known to interact genetically with sdc-1 and sdc-3 to implement sex determination and dosage compensation in XXanimals (Villeneuve and Meyer 1990 Davis and Meyer 1997 Dawes et al. 1999), but the precise molecular roles of SDC-1 and SDC-3 were not understood. Here we present three lines of evidence that the three SDC proteins form a complex in vivo to repress her-1 andX chromosomes directly.
First, SDC-1, SDC-2, and SDC-3 all colocalize to X chromosomes and to her-1 regulatory regions in vivo. Hermaphrodites carrying multiple tandem copies of her-1 regulatory regions on GFP-tagged extrachromosomal arrays were stained with affinity-purified SDC antibodies (see Materials and Methods). SDC protein localization was assessed in adult intestinal nuclei, whose large size and polyploid DNA content facilitate the assay. The highly charged SDC-2 protein, which bears a coiled-coil motif, localized to X chromosomes and her-1 arrays in adult gut cells (Fig.1A), as shown previously in embryos (Dawes et al. 1999), thus validating the assay. The zinc-finger proteins SDC-1 and SDC-3 (Nonet and Meyer 1991 Klein and Meyer 1993) colocalized with SDC-2 at her-1 and on X chromosomes (Fig. 1A,B), consistent with a direct role for these proteins in her-1repression and dosage compensation. In XX animals carrying ansdc-3(Tra) mutation, localization of SDC-1, SDC-2, and SDC-3 to her-1 was greatly reduced, but localization to theX chromosome appeared unaffected (Fig. 1F data not shown), consistent with the mutation impairing sex determination but not dosage compensation (DeLong et al. 1993). sdc-3(Tra) derepressesher-1 transcription, causing 100% of XX animals to be severely masculinized by disrupting a putative ATP-binding motif in SDC-3 (DeLong et al. 1993 Klein and Meyer 1993). Therefore, SDC-1, SDC-2, and SDC-3 are localized appropriately to achieve both gene-specific and chromosome-wide repression.
The X-chromosome dosage compensation machinery localizes to her-1 regulatory regions in vivo. Confocal images of an individual gut nucleus (A–F) or an embryonic nucleus (G,H) from wild-type or mutant [sdc-3(Tra) or dpy-27] XX animals immunostained with SDC, DPY, or MIX antibodies, as indicated in each panel. The nuclei contain extrachromosomal DNA arrays carrying multiple copies of her-1 regulatory regions (plasmid pHD25 of Fig. 3A),lac operator repeats (lacO), and a transgene encoding a LacI–GFP fusion protein. LacI–GFP repressor binding tolacO permits array detection by GFP autofluorescence. Colocalization (yellow) between arrays (green) and antibodies (red) in the merged images (right panels) showed association of the protein with her-1 regulatory sequences. Arrowheads mark theX chromosomes. Consistent with sdc-3(Tra) causing derepression of her-1, it blocks SDC and DPY proteins from associating with her-1.
Second, the SDC proteins interact physically to form a complex. Antibodies to any one of the SDC proteins coimmunoprecipitated all three SDC proteins from wild-type embryonic extracts (Fig.2B). These precipitation reactions were specific, because none of the preimmune sera precipitated any of the SDC proteins (Fig. 2B), and none of the SDC antibodies precipitated (data not shown) or identified (Fig. 2A) their cognate proteins from extracts of the respective sdc null mutants.
The SDC proteins form a complex in vivo. (A) Detection of SDC proteins in embryonic extracts. Western blots of extracts from wild-type (N2) or sdc (null) mutant embryos carrying a deletion or nonsense mutation in the sdc gene were probed with the SDC antibody indicated on the left. Proteins of 250 kD, 240 kD (a doublet), and 140 kD were detected by SDC-2, SDC-3, or SDC-1 antibodies, respectively, in wild-type but not sdc (null) extracts, showing antibody specificity. (B) Antibody to any one SDC protein coimmunoprecipitated all three SDC proteins. Coimmunoprecipitations were performed with each SDC antibody on wild-type embryonic extracts. The coimmunoprecipitated material was separated by SDS-PAGE and immunoblotted with the antibody indicated on the left. (PI) Preimmune sera for SDC-1 antibody, (IP) immunoprecipitation.
Third, the SDC complex represses transcription of her-1, since ectopic production of SDC proteins in XO animals (normally males) induced hermaphrodite sexual development. SDC-2 is normally expressed only in XX animals, and ectopic expression of SDC-2 transformed 36% of XO animals into hermaphrodites (Dawes et al. 1999), a sexual transformation that required wild-typesdc-3 activity. Given the incomplete feminization with SDC-2 alone, we simultaneously overexpressed SDC-2 with either SDC-1 or SDC-3 to assess their combined contributions toward hermaphrodite development. Overexpression of only SDC-1 (data not shown) or SDC-3 (Davis and Meyer 1997) failed to feminize XO animals. However, overexpression of both SDC-2 and SDC-1 greatly enhanced the XOfeminization, causing ∼88% of these XO animals to be sexually transformed (Table 1). Nearly all were self-fertile, in contrast to transformed XO animals that expressed only SDC-2. Increasing the level of SDC-3 in XOanimals, as verified by antibody staining, did not enhance the feminization caused by SDC-2 (31% feminization with both proteins), indicating that SDC-3 was not limiting (Table 1). The synergy between SDC-1 and SDC-2 in feminizing XO animals in an SDC-3-dependent manner provides functional evidence that all three SDC proteins act together to repress her-1 directly.
sdc-1 enhances sdc-2 in feminizingXO animals
SDC proteins recruit the X-chromosome dosage compensation complex to her-1
Because SDC-2 and SDC-3 play pivotal roles in assembling the dosage compensation complex onto X chromosomes (Chuang et al. 1996Davis and Meyer 1997 Dawes et al. 1999), we asked whether SDC proteins recruit this complex to her-1. The dosage compensation complex includes the dosage compensation-specific protein DPY-27 and the dual-function proteins DPY-26 and MIX-1, which also act in meiosis and mitosis, respectively (Chuang et al. 1996 Lieb et al. 1996, 1998). The dosage compensation proteins resemble components of the widely conserved condensin complex, which drives mitotic chromosome condensation in vitro, implying that regulation ofX-chromosome expression involves modulation of chromatin structure (Koshland and Strunnikov 1996 Hirano 2000). All dosage compensation proteins except SDC-2 require SDC-3 for their localization to the X chromosome (Chuang et al. 1996 Davis and Meyer 1997), and SDC-3, in turn, requires SDC-2 for its localization to theX chromosome (Davis and Meyer 1997). SDC-2 can localize to theX chromosome without other dosage compensation proteins (Dawes et al. 1999), suggesting that it recognizes the X chromosome and confers chromosome specificity to dosage compensation. Not all of the dosage compensation components appear essential for her-1repression, because the rare dpy-26, dpy-27, ordpy-28 XX mutants that escape lethality develop as hermaphrodites (Plenefisch et al. 1989). Therefore, discovery of the complete dosage compensation machinery on her-1 would show that SDC-2 targets this machinery to the chromatin it binds.
DPY-26, DPY-27, and MIX-1 all colocalized with SDC proteins on bothher-1 arrays and X chromosomes (Fig. 1C–E). Furthermore, localization of the three dosage compensation proteins toher-1 was dependent on SDC proteins because thesdc-3(Tra) mutation disrupted the localization toher-1 but not to the X chromosome (Fig. 1F data not shown). The localization of SDC-2 and SDC-3 was not dependent on DPY-27 (Fig. 1G), but DPY-26 required DPY-27 for its localization toher-1 (Fig. 1H), as it does for its localization to theX chromosome. Thus, assembly of known dosage compensation components onto her-1 resembles their assembly onto theX chromosome. Moreover, SDC-2 recruits the dosage compensation machinery to its chromatin targets, even though some components may be dispensable for repression.
The SDC/dosage compensation complex associates with three different chromatin targets within her-1
We determined the exact sites within her-1 that recruit the dosage compensation machinery. Using the extrachromosomal array assay to examine individual 1-kb fragments across her-1, we found that SDC-1, SDC-2, SDC-3, DPY-26, DPY-27, and MIX-1 all colocalized with three different regions defined by fragments B, C, and D (Fig.3A,B). Localization of all proteins was disrupted by an sdc-3(Tra) mutation (data not shown), showing that binding was specific.
SDC-2 localization to her-1 is specified by three distinct DNA recognition elements whose binding capacity is disrupted by specific mutations. (A) Schematic of theher-1 gene and summary of subregions tested for SDC-2 colocalization by the array assay. Transcription from the P1 promoter produces the functional male-specific her-1 transcript, including four exons (green). A promoter (P2) resides within the second intron of her-1. P2 is coregulated with P1 and makes a 0.8-kb transcript of unknown function that includes the last two exons ofher-1 (Trent et al. 1991 Perry et al. 1993). The degree of SDC-2 colocalization with her-1 regions is shown by color, with the key on the right. The three smallest regions with strong SDC-2 colocalization (region 1 [B], region 2 [C5], and region 3 [D6]) are shown by dark gray shading. C5 and D6 share an identical 15-bp element (solid vertical lines) and 50% overall sequence identity. B has no obvious similarity with C5 or D5 but contains the site ofher-1(gf) (dashed vertical line). SDC-2 colocalization was completely disrupted by a randomized 15-mer in either C5 or D5 and by the G → A transition of her-1(gf) in B (yellow stars in B‘, C5′, and D5′). (B) Confocal images of an individual gut nucleus from a wild-type XX animal bearing GFP-tagged extrachromosomal arrays (green) with either wild-type (B, C5, and D5) or mutant (B‘, C5′, and D5′) versions of regions 1–3. Animals were immunostained with antibodies to DPY-27 (red) or SDC-3 (blue). Colocalization between the array and protein appears as yellow in the merged image. Arrowheads indicate X chromosomes.
Fragment B includes the P1 promoter that produces the 1.2-kb functionalher-1 transcript (Trent et al. 1991 Perry et al. 1993). This regulatory region was first implicated in her-1 repression by a gain-of-function mutation, her-1(gf), that partially derepresses her-1 transcription, causing substantial, but incomplete masculinization of XX mutants (Trent et al. 1988). The location of her-1(gf) 2 bp before the transcriptional start site prompted us to test whether her-1(gf) interferes with binding of the repression complex (Perry et al. 1994). Indeed, SDC-2, SDC-3, and DPY-27 failed to associate with a fragment (B‘) harboring the A → T transition of her-1(gf), showing that derepression of her-1 transcription is caused at least in part by disrupting repressor binding (Fig. 3A,B data not shown). Theher-1(gf) mutation appears to eliminate rather than reduce SDC binding, because overexpression of SDC-2 failed to suppress theXX masculinization caused by her-1(gf) (see Materials and Methods).
Fragments C and D are within the large second intron of her-1. Unlike P1, this specific region had not been implicated inher-1 repression by gain-of-function mutations. However, the partial masculinization of XX animals by her-1(gf) compared with the nearly complete masculinization bysdc-3(Tra) suggested that SDC-mediated her-1repression requires sequences outside the gf region. Moreover, indirect experiments suggested a possible involvement of the second intron in her-1 repression (Li et al. 1999).
We explored whether the protein–DNA interactions observed with fragments C and D on extrachromosomal arrays also occurred at the endogenous her-1 gene by performing chromatin immunoprecipitations (ChIP) from lysates of formaldehyde-treated wild-type embryos. SDC-2 antibodies were used to immunoprecipitate the SDC complex with its associated DNA, and the DNA was analyzed for enrichment of her-1 fragments using primers to regions A–F in separate PCR reactions. Primers flanking him-1, a gene on a different chromosome, and genomic fragments just upstream of region A were used as controls. Only DNA from regions C and D was specifically enriched by threefold to fourfold relative to the negative control (Fig. 4A). In parallel ChIPs performed with SDC-2 or SDC-3 antibodies, only DNA from wild-type, but notsdc-3(Tra) lysates was enriched for fragment C (Fig. 4B). This result confirmed the specificity of the ChIP by showing that it correctly reflects the disruption of SDC binding to her-1caused by sdc-3(Tra). In controls, equivalent levels of region C DNA were detected in PCRs using DNA extracted from wild-type and mutant lysates (Fig. 4B). Likewise, comparable levels of SDC proteins were detected in both lysates with Western blots (Fig. 4C) and IP experiments (Fig. 4D). Together these experiments show that the SDC complex associates with regions C and D in the endogenousher-1 gene. The inability to detect region B by this assay suggested that region B has a lower capacity for SDC binding than regions C and D, as shown below and hypothesized previously (Li et al. 1999).
Chromatin immunoprecipitation experiments (ChIP) show the interaction of SDC proteins with the endogenous her-1gene. (A) PCR analysis of DNA from a ChIP performed with SDC-2 antibodies and lysates of formaldehyde cross-linked XXembryos. Primer sets specific to her-1 regions or a control gene (him-1) were used for PCRs with mock-precipitated DNA (M) and twofold serial dilutions of SDC-2-precipitated DNA (SDC-2) or input DNA (Input). The intensity of the PCR band produced by each primer pair from IP-enriched DNA was normalized to the corresponding PCR band produced from the highest concentration of input DNA. Regions D and C of her-1 were specifically enriched above him-1control DNA by threefold or fourfold, respectively. (Primers flankinghim-1 produced a PCR product from IP-enriched DNA of 22% normalized intensity, whereas primers flanking regions D and C produced bands of 67% and 86%, respectively, normalized intensity.) (B) PCR analysis with region C primers was performed on twofold serial dilutions of DNA from a ChIP using SDC-2 or SDC-3 antibodies and lysates of formaldehyde-treated wild-type orsdc-3(Tra) XX embryos. The intensity of each PCR band was normalized to the intensity of the PCR band made from the highest concentration of IP-enriched DNA from the wild-type lysate. The average intensities and standard deviations were calculated from four sets of PCR analyses on material from two independent ChIP experiments. The specificity of the ChIP protocol was shown by the precipitation of region C DNA from wild-type but not mutant lysates. Similar levels of region C DNA were detected by PCR using twofold serial dilutions of wild-type and sdc-3(Tra) input lysates. (C,D) Similar levels of SDC-2 and SDC-3 were detected by Western blot analysis of either (C) whole lysates or (D) SDC-2 IP material from lysates of formaldehyde-treated wild-type and sdc-3(Tra) embryos.
Diverse DNA recognition elements reside within the three distincther-1 chromatin targets
Having shown that fragments B, C, and D are true SDC targets, we defined the DNA sequence requirements for SDC binding more precisely. Of four region B subfragments, only B2 supported significant SDC-2 colocalization (Fig. 3A). However, localization to B2 was less consistent than localization to B, suggesting that strong SDC-2 binding requires more than one discrete element within region B. In contrast, the robust localization to regions C and D was narrowed to a 303-bp fragment (C5) of C and to a 192-bp fragment (D6) of D (Fig. 3A,B). SDC-1, SDC-3, DPY-26, DPY-27, and MIX-1 all colocalized with SDC-2 on these fragments, showing that all the information required for SDC proteins to interact with chromatin and recruit the dosage compensation complex can be specified by 192 bp of DNA (Fig. 3A,B data not shown) that has been removed from its native chromosomal context.
Very limited similarity in DNA sequences was found between B (region 1) and either C5 (region 2) or D6 (region 3). In contrast, C5 and D5 (a 287-bp fragment that includes D6) share 50% overall identity and a 15-bp stretch (CAAAAACTGAGCCTG) of complete identity on the antisense strand of C5 and the sense strand of D5. An exact copy of this element is not found on X or elsewhere in the genome. Randomizing the 15-bp element to ACAGACTGCAGATAC (for C5′ and D5′) or GA CAGACGTCAATAC (for D5′) prevented SDC-2, SDC-3, and DPY-27 proteins from localizing to arrays with the mutant fragments (Fig.3A,B data not shown). The 15-bp repeated sequence is therefore necessary for targeting SDC and DPY proteins to regions 2 and 3. The 15-mer is not sufficient, however, because SDC-2 failed to associate with arrays carrying multiple copies of random DNA and either the 15-mer or a 28-bp element that includes the 15-mer and neighboring common sequences (data not shown). Other cis-acting sequences must be essential. Thus, the three DNA elements used to target the dosage compensation machinery to her-1 are diverse. Either SDC proteins themselves have flexibility in sequence recognition or other cellular components help confer sequence specificity and binding.
Differential recruitment of SDC-2 to her-1 by individual recognition elements
Identification of mutations that eliminated SDC binding to individual sites in her-1 allowed us to assess and correlate in vivo the functional contribution of each site toward overall SDC binding and repression of her-1. We introduced theher-1(gf) mutation of region 1 and the randomized 15-mers of regions 2 and 3 together or separately into full-lengthher-1-rescuing constructs to test the role of each site.XX animals expressing a single construct from a GFP-tagged extrachromosomal array were examined for the frequency of SDC-2 localization to arrays and for sexual transformation to the male fate, an indicator of transcriptional derepression (see Materials and Methods). The functional significance of a site could then be inferred by comparing the change in SDC localization with the degree of sexual transformation caused by disrupting that site (Fig.5A).
The relative contributions of the three her-1recognition elements differ for SDC binding and her-1repression. (A) Mutation of specific DNA sequences within full-length her-1 transgenes disrupts SDC-2 localization toher-1 and repression of her-1. her-1constructs included in transgenic arrays are depicted by diagrams on the left, with mutations indicated by white stars. The percent SDC-2 localization to transgenic arrays is represented by gray bars. The standard deviation between lines assayed is represented by a dotted line. (n) Total number of nuclei scored in all lines. Masculinization caused by the full-length her-1 transgenes was quantified (see Materials and Methods) and then rated as (−) none, (+) weak, (++) moderate, (+++) strong, or (++++) severe. (B–G) Examples of masculinization caused by derepression of her-1. DIC photomicrographs of tails from (B) a wild-type XX hermaphrodite, (C–F) XX animals masculinized by full-length her-1 transgenes, and (G) a wild-typeXO male. Lateral views (B–D) and ventral views (E–G). (White arrow) male fan (black arrow) male sensory rays (black arrowhead) male spicules.
Regions 2 and 3 had approximately equivalent SDC-2 binding activity in the context of the full-length her-1 gene, and these regions supported more robust binding than region 1 (Fig. 5A). SDC-2 localized to 90% of arrays carrying a wild-type her-1 gene. The localization was reduced only slightly, to 85%, by theher-1(gf) lesion in region 1, even though this lesion abolished SDC-2 localization to a fragment containing only region 1. The remaining SDC binding must have occurred through regions 2 and 3. Indeed, in transgenes with a wild-type region 1, randomization of either 15-mer decreased SDC-2 localization to 20%–40%, and randomization of both 15-mers decreased SDC-2 localization to 10%. SDC localization was not significantly reduced by disrupting region 1 on a transgene already mutant for either region 2 or region 3, but was mildly reduced by disrupting region 1 on a transgene mutant for both regions 2 and 3. The comparatively weak SDC binding affinity for region 1 in the context of the full-length her-1 transgene is consistent with the difficulty in detecting region 1 by ChIP analysis on the endogenous gene.
Complete repression of her-1 requires the participation of all three SDC-binding regions
Is the strength of SDC binding to a region correlated with effectiveness in repressing her-1? Analysis of sexual phenotype in XX animals with wild-type or modified full-lengthher-1 transgenes revealed that complete repression ofher-1 required the participation of all three SDC-binding regions. However, region 1, the weakest in SDC-binding activity, made the greatest single contribution to repression (Fig. 5A–G). Regions 2 and 3 contributed repression activity in the absence of region 1, but repression was less effective than from region 1 (Fig. 5A–G). The degree of her-1 repression from regions 2 and 3 may be more comparable to the repression of X-linked genes that occurs during dosage compensation.
These conclusions were drawn from the following observations (Fig.5A–G): XX animals carrying wild-type her-1transgenes showed very low levels (−) of masculinization, indicating strong repression. Mutation of either region 2 or 3 caused weak masculinization (+) that was correlated with intermediate disruption of SDC-2 localization. Mutation of both regions 2 and 3 caused moderate masculinization (++), despite causing strong disruption of SDC-2 localization. Finally, mutation of region 1 caused strong masculinization (+++), despite causing only a slight reduction in SDC localization. This masculinization was not enhanced by disrupting only region 2 or 3 but was enhanced by disrupting both regions 2 and 3, causing severe masculinization (++++).
The weak binding of SDC proteins to region 1 on a full-lengthher-1 transgene raised the concern that SDC proteins may not mediate repression from region 1. Therefore, we assessed the impact of disrupting SDC binding specifically to region 1 by introducing ansdc-3(Tra) mutation into genotypically her-1(−) animals carrying full-length her-1(+) transgenes with mutations in regions 2 and 3. A twofold to threefold increase in the number of masculinized animals was found, showing that SDC proteins contribute to repression from region 1 in vivo.
The involvement of regions 2 and 3 in SDC-mediated her-1repression in vivo is further reinforced by interpreting previous genetic data in the context of the SDC-binding data. Our results show that the her-1(gf) mutation eliminates SDC binding to region 1 but not to regions 2 and 3, whereas the sdc-3(Tra) mutation severely reduces SDC binding to all three regions. Thus, the greater degree of masculinization in XX animals caused by thesdc-3(Tra) mutation (100% of mutants) compared with theher-1(gf) mutation (30% of mutants) must result from reduction of SDC binding to regions 2 and 3. Therefore, regions 2 and 3 contribute substantially to SDC-mediated repression of the endogenousher-1 gene, along with region 1.
Classification and origin of eccDNAs
The genome of eukaryotes consists of chromosomal DNA and extrachromosomal DNA elements which are physically excised from the chromosomes 3 . The term NA” is now used to describe the full spectrum of circular DNAs in eukaryotes. eccDNAs are widely spread across various eukaryotes from yeast to human 12 - 14 . eccDNAs can be divided into organelle eccDNAs such as mitochondrial DNAs (mtDNAs), and more flexible non-organelle eccDNAs such as double minutes (DMs), episomes, small polydispersed circular DNAs (spcDNAs) and microDNAs (Table (Table1) 1 ) 15 . Their sizes range from hundreds of base pairs (bp) to as large as several mega bases (Mb). Owing to universal existence and heterogeneous derivation, eccDNAs are considered to reflect genomic plasticity and instability.
Classification of eccDNAs in eukaryotes
|Name of the eccDNA||Size range||Biological function||References|
|Mitochondrial DNA||16 kb||Maintaining mitochondria function||134|
|Double minute||100 kb-3 Mb||Acting as a vehicle for extrachromosomal gene amplification||6, 56|
|Small polydispersed circular DNA||100 bp-10 kb||Enhancing genomic instability||34|
|microDNA||100-400 bp||Producing miRNAs||75|
|Telomeric circle||Integral multiples of 738 bp||Restoring telomere length||135|
Increasing evidence indicates that eccDNAs in eukaryotic cells usually carry interspersed repeat sequences or tandemly repeated genomic sequences 16 - 18 . It can be concluded that tandemly repetitive DNAs are particularly prone to eccDNA formation 17 . On the other hand, eccDNAs also derive from nonrepetitive DNA. Loon et al. 19 found that eccDNAs from HeLa S3 cells consisted of nonrepetitive or low-copy DNA sequences. Plus, some nonrepetitive spcDNAs were reported to be bordered on both sides by direct repeats of a mean length of 9-11 bp 20 , 21 . eccDNAs can originate from both coding and noncoding regions. For instance, oncogenes and drug resistance genes have been identified as predominant components of DMs 22 . In addition, microDNAs are preferentially stemmed from exons, 5' untranslational regions (5' UTR) and CpG islands 23 .
NIPT Technology Types
The technology platforms commonly used for NIPT are whole-genome sequencing using next-generation sequencing (NGS) or various targeted methods. When looking at the best NIPT technology fit for your lab, you’ll want to consider data generation and analysis, lab workflow, and resulting clinical implications.
NIPT and Whole-Genome Sequencing
NIPT using whole-genome sequencing technology provides the most informative NIPT results 1-7 with a comprehensive view across the entire genome. NIPT with whole-genome sequencing consistently has lower failure rates compared to targeted sequencing or array-based platforms 8 .
The polymerase chain reaction (PCR)-free sample preparation used with whole-genome-sequencing-based NIPT improves laboratory workflow, greatly reduces assay complexity and significantly improves turn-around time (TAT). This type of NIPT NGS technology can also be scaled easily to accommodate the needs of a growing lab.
NIPT and Whole-Genome Sequencing
Targeted Approaches for NIPT
Targeted technologies for NIPT include single nucleotide polymorphism (SNP) analysis, microarray analysis, and rolling circle amplification. With targeted approaches, only limited regions of select chromosomes are analyzed.
These targeted approaches have additional steps and employ more rounds of amplification than whole-genome sequencing methods, introducing a more complicated workflow.
SNPs are genetic variations among individuals. This technique determines the difference between parent and child DNA, and the relative dosage of genetic variation to infer copy number. cfDNA is amplified by PCR using specific SNP targets. These targets are sequenced, and the allele distribution of both mother and child are determined. An algorithm determines abnormalities in expected allele frequencies.
In this type of NIPT, a microarray is used, which is an assembly of microscopic DNA regions attached to a solid surface. cfDNA fragments are amplified by PCR, tagged with a fluorescent probe, and bind to complimentary sequences on the NIPT microarray. Both the light intensity and binding position indicate the relative amount of DNA and presence or absence of the target, respectively. Deviations in expected fluorescent counts indicate aneuploidy.
Rolling Circle Amplification
Rolling circle amplification targets specific cfDNA fragments which bind to a circular template and replicate by a rolling mechanism. The rolling circle replication products are fluorescently labeled and counted. Deviations in expected fluorescent counts indicate aneuploidy.
Rolling Circle Amplification
A Guide to NIPT Technology Options
Learn more about evaluating NIPT options and key considerations when selecting a technology for your lab.
A Guide to NIPT Technology Options
Comparison of NIPT Technology Types
How is NIPT Data Generated?
Purified cfDNA fragments are labelled with universal sequencing adapters to create a library. The library is amplified through bridge amplification (PCR-free) or PCR.
The sample libraries are sequenced using single or paired-end sequencing.
SNPs on chromosomes of interest are amplified by PCR and sequenced from cfDNA isolated from the plasma of pregnant women.
Probes designed to match specific sequences of the genome are synthesized onto specific areas on the microarray.
Targeted cfDNA fragments are amplified by PCR, tagged with a fluorescent probe, and bind to complimentary sequences on the microarray.
Target cfDNA fragments hybridize to probes designed to form circular complexes.
The DNA circles are copied thousands of times to generate rolling circle products.
How is NIPT Data Analyzed?
An over or under representation of sequenced reads based on expected distribution indicates aneuploidy.
An altered ratio of target alleles compared with expected ratios indicates aneuploidy.
An over or under representation of fluorescent counts compared with a reference genome indicates aneuploidy.
An over or under representation of fluorescent counts compared with a reference genome indicates aneuploidy.
What are the Clinical Considerations of NIPT?
All chromosomes are evaluated.
Only select chromosomes are evaluated.
Only select chromosomes are evaluated.
Only select chromosomes are evaluated.
What are the NIPT Lab Workflow Implications?
A PCR-free workflow eliminates the need for pre- and post-PCR lab space, allowing steps to be performed in a single area.
This workflow requires less hands-on time, faster TAT, and is less prone to contamination since PCR amplification is not employed.
PCR involves a more complicated workflow, significantly longer TAT, and necessitates pre- and post-PCR workspace considerations.
PCR involves a more complicated workflow, significantly longer TAT, and necessitates pre- and post-PCR workspace considerations.
There is reduced risk for contamination since PCR is not employed however, this method results in a long TAT.
The Science Behind NIPT
See how this NIPT workflow detects aneuploidy in three simple steps.
Noninvasive Prenatal Testing and NIPT
Dr. Wapner, Vice Chair of Research in Obstetrics and Gynecology for Columbia University, shares his perspective on prenatal screening and NIPT.
NIPT Delivers Relief to Expectant Mother
Whole-genome sequencing provides results when another test was inconclusive.
NIPT: Effective Screening for the General Pregnancy Population
Glenn Palomaki, PhD, discusses the clinical validity and utility of offering NIPT as a primary screen in the general pregnancy population.
- Illumina Inc. VeriSeq™ NIPT Solution v2 Package Insert. 2020
- Palomaki GE, Kloza EM, Lambert-Messerlian GM, et al. DNA sequencing of maternal plasma to detect Down syndrome: an international clinical validation study. Genet Med. 201113(11):913-920
- Ryan A, Hunkapiller N, Banjevic M, et al.Validation of an Enhanced Version of a Single-Nucleotide Polymorphism-Based Noninvasive Prenatal Test for Detection of Fetal Aneuploidies. Fetal Diagn Ther. 201640(3):219-223.
- Stokowski R, Wang E, White K, et al. Clinical performance of non-invasive prenatal testing (NIPT) using targeted cell-free DNA analysis in maternal plasma with microarrays or next generation sequencing (NGS) is consistent across multiple controlled clinical studies. Prenat Diagn. 2015doi: 10.1002/pd.4686.
- Jones KJ, Wang E, Bogard P, et al. Targeted cell-free DNA analysis with microarray quantitation for assessment of fetal sex and sex chromosome aneuploidy risk. Ultrasound Obstet Gynecol. 201851(2):275-276.
- Taneja PA, Snyder HL, de Feo E, et al. Noninvasive prenatal testing in the general obstetric population: clinical performance and counseling considerations in over 85,000 cases. Prenat Diagn. 2016 doi:10.1002/pd.4766.
- McCullough RM, Almasri EA, Guan X, et al. Non-invasive prenatal chromosomal aneuploidy testing--clinical experience: 100,000 clinical samples.
- Data on file. Illumina, Inc. November 2018.
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Although most DNA is contained within a cell’s chromosomes, many cells have additional molecules of DNA outside the chromosomes, called extrachromosomal DNA, that are also part of its genome. The genomes of eukaryotic cells would also include the chromosomes from any organelles such as mitochondria and/or chloroplasts that these cells maintain (Figure 3). The maintenance of circular chromosomes in these organelles is a vestige of their prokaryotic origins and supports the endosymbiotic theory (see Foundations of Modern Cell Theory). In some cases, genomes of certain DNA viruses can also be maintained independently in host cells during latent viral infection. In these cases, these viruses are another form of extrachromosomal DNA. For example, the human papillomavirus (HPV) may be maintained in infected cells in this way.
Figure 3. The genome of a eukaryotic cell consists of the chromosome housed in the nucleus, and extrachromosomal DNA found in the mitochondria (all cells) and chloroplasts (plants and algae).
Besides chromosomes, some prokaryotes also have smaller loops of DNA called plasmids that may contain one or a few genes not essential for normal growth (see Figure 1 in Unique Characteristics of Prokaryotic Cells). Bacteria can exchange these plasmids with other bacteria in a process known as horizontal gene transfer (HGT). The exchange of genetic material on plasmids sometimes provides microbes with new genes beneficial for growth and survival under special conditions. In some cases, genes obtained from plasmids may have clinical implications, encoding virulence factors that give a microbe the ability to cause disease or make a microbe resistant to certain antibiotics. Plasmids are also used heavily in genetic engineering and biotechnology as a way to move genes from one cell to another. The role of plasmids in horizontal gene transfer and biotechnology will be discussed further in Mechanisms of Microbial Genetics and Modern Applications of Microbial Genetics.
Think about It
Maria, a 20-year-old anthropology student from Texas, recently became ill in the African nation of Botswana, where she was conducting research as part of a study-abroad program. Maria’s research was focused on traditional African methods of tanning hides for the production of leather. Over a period of three weeks, she visited a tannery daily for several hours to observe and participate in the tanning process. One day, after returning from the tannery, Maria developed a fever, chills, and a headache, along with chest pain, muscle aches, nausea, and other flu-like symptoms. Initially, she was not concerned, but when her fever spiked and she began to cough up blood, her African host family became alarmed and rushed her to the hospital, where her condition continued to worsen.
After learning about her recent work at the tannery, the physician suspected that Maria had been exposed to anthrax. He ordered a chest X-ray, a blood sample, and a spinal tap, and immediately started her on a course of intravenous penicillin. Unfortunately, lab tests confirmed the physician’s presumptive diagnosis. Maria’s chest X-ray exhibited pleural effusion, the accumulation of fluid in the space between the pleural membranes, and a Gram stain of her blood revealed the presence of gram-positive, rod-shaped bacteria in short chains, consistent with Bacillus anthracis. Blood and bacteria were also shown to be present in her cerebrospinal fluid, indicating that the infection had progressed to meningitis. Despite supportive treatment and aggressive antibiotic therapy, Maria slipped into an unresponsive state and died three days later.
Anthrax is a disease caused by the introduction of endospores from the gram-positive bacterium B. anthracis into the body. Once infected, patients typically develop meningitis, often with fatal results. In Maria’s case, she inhaled the endospores while handling the hides of animals that had been infected.
The genome of B. anthracis illustrates how small structural differences can lead to major differences in virulence. In 2003, the genomes of B. anthracis and Bacillus cereus, a similar but less pathogenic bacterium of the same genus, were sequenced and compared.  Researchers discovered that the 16S rRNA gene sequences of these bacteria are more than 99% identical, meaning that they are actually members of the same species despite their traditional classification as separate species. Although their chromosomal sequences also revealed a great deal of similarity, several virulence factors of B. anthracis were found to be encoded on two large plasmids not found in B. cereus. The plasmid pX01 encodes a three-part toxin that suppresses the host immune system, whereas the plasmid pX02 encodes a capsular polysaccharide that further protects the bacterium from the host immune system (Figure 4). Since B. cereus lacks these plasmids, it does not produce these virulence factors, and although it is still pathogenic, it is typically associated with mild cases of diarrhea from which the body can quickly recover. Unfortunately for Maria, the presence of these toxin-encoding plasmids in B. anthracis gives it its lethal virulence.
Figure 4. Genome sequencing of Bacillus anthracis and its close relative B. cereus reveals that the pathogenicity of B. anthracis is due to the maintenance of two plasmids, pX01 and pX02, which encode virulence factors.
- What do you think would happen to the pathogenicity of B. anthracis if it lost one or both of its plasmids?
Clinical Focus: Aamir, Resolution
Within 24 hours, the results of the diagnostic test analysis of Aamir’s stool sample revealed that it was positive for heat-labile enterotoxin (LT), heat-stabile enterotoxin (ST), and colonization factor (CF), confirming the hospital physician’s suspicion of ETEC. During a follow-up with Aamir’s family physician, this physician noted that Aamir’s symptoms were not resolving quickly and he was experiencing discomfort that was preventing him from returning to classes. The family physician prescribed Aamir a course of ciprofloxacin to resolve his symptoms. Fortunately, the ciprofloxacin resolved Aamir’s symptoms within a few days.
Aamir likely got his infection from ingesting contaminated food or water. Emerging industrialized countries like Mexico are still developing sanitation practices that prevent the contamination of water with fecal material. Travelers in such countries should avoid the ingestion of undercooked foods, especially meats, seafood, vegetables, and unpasteurized dairy products. They should also avoid use of water that has not been treated this includes drinking water, ice cubes, and even water used for brushing teeth. Using bottled water for these purposes is a good alternative. Good hygiene (handwashing) can also aid the prevention of an ETEC infection. Aamir had not been careful about his food or water consumption, which led to his illness.
Aamir’s symptoms were very similar to those of cholera, caused by the gram-negative bacterium Vibrio cholerae, which also produces a toxin similar to ST and LT. At some point in the evolutionary history of ETEC, a nonpathogenic strain of E. coli similar to those typically found in the gut may have acquired the genes encoding the ST and LT toxins from V. cholerae. The fact that the genes encoding those toxins are encoded on extrachromosomal plasmids in ETEC supports the idea that these genes were acquired by E. coli and are likely maintained in bacterial populations through horizontal gene transfer.
In both Drosophila and C. elegans, global silencing mechanisms appear to play a crucial role in the specification and maintenance of germ line tissue (Wylie, 1999). In C. elegans, these silencing mechanisms can be studied using transgene arrays containing a GFP reporter gene under the control of a promoter normally expressed in all cells (Kelly et al., 1997). The transgene arrays are strongly silenced in germ cells but are reactivated in the soma of each generation. Silencing in the germline can be partially prevented by increasing the ‘complexity’ of the transgene arrays formed in vivo through the co-injection of excess amounts of random, linear genomic fragments (Kelly et al., 1997).
Germline silencing of transgene arrays requires the action of the MES proteins (maternal effect sterility), MES-2, -3, -4, and -6 (Kelly and Fire, 1998). Two of the mes genes, mes-2 and mes-6, encode worm homologs of the Drosophila Polycomb Group proteins, Enhancer of Zeste and Extra Sex Combs, respectively (Holdeman et al., 1998 Korf et al., 1998). The Polycomb Group proteins maintain transcriptional repression of developmentally regulated genes through their ability to modulate chromatin conformation (Kennison, 1995). In addition, MES-2 and MES-4 each contain a SET domain, a conserved feature of many chromatin-interacting proteins. Defects in the mes factors cause sterility, owing to germ cell degeneration, and alleviate silencing of transgene arrays in the germline. Similar phenotypes can also result from depletion of a histone H1 isoform, H1.1 (Jedrusik and Schulze, 2001). Gene silencing through the regulation of chromatin conformation is therefore likely to be an essential component of germline maintenance, but the endogenous targets of this regulation are poorly understood. Severity of the germ cell degeneration in mes mutant animals increases with X-chromosome dose (Garvin et al., 1998), suggesting that some of the gene targets of MES-induced silencing reside on the X chromosome.
Global gene expression analysis has also suggested that the X chromosome is a possible target of silencing in the C. elegans germline. Microarray analyses have identified 1416 germline-enriched genes in C. elegans, which were classified into three distinct groups: sperm-enriched, oocyte-enriched and germline-intrinsic genes (defined as genes expressed similarly in the germline regardless of the gamete being made) (Reinke et al., 2000). Strikingly, sperm-enriched and germline-intrinsic genes are almost completely absent from the X chromosome. By contrast, oocyte-enriched genes are present on the X chromosome at a similar frequency to those found on autosomes. C. elegans XO males make only sperm and thus would not require expression of oocyte-enriched genes, whereas XX hermaphrodites first produce sperm as L4 larvae and then become strictly oogenic as adults (Schedl, 1997 Hubbard and Greenstein, 2000). The X chromosome in male germlines may therefore be regulated differently from autosomes in a manner that prohibits the presence of the sperm-enriched and germline-intrinsic genes. Another possibility is that these classes of genes are absent from the X chromosome for some unknown reason, but that the X chromosome is otherwise competent for gene expression.
In support of the first possibility, the distinction of the male X chromosome from the autosomes in the germline of C. elegans is reminiscent of sex chromatin formation in other species that bear non-equivalent sex chromosomes (heterogametic: XO or XY). During the pachytene stage of C. elegans meiosis in males, the single (and thus unpaired) X chromosome adopts a highly compact morphology analogous to that seen in mammalian spermatocytes (Goldstein, 1982). In the heterogametic sex of diverse species, the male X chromosome in the pachytene stage of meiotic prophase is found in a visually distinct structure called the XY- or sex-body that is transcriptionally inactive (Handel and Hunt, 1992). McKee and Handel (McKee and Handel, 1993) proposed that the condensation of sex chromatin in XY and XO male germlines, and by consequence the transcriptional inactivation of these chromosomes, prevents harmful recombination events between non-equivalent X and Y chromosomes, and prevents loss of a single chromosome lacking a pairing partner in XO animals. In C. elegans, both the exclusion of sperm-enriched and germline-intrinsic genes from the X chromosome, and the condensed structure of the X chromosome in the XO male germline suggest that the X chromosome in the male germ line may be targeted for silencing.
If the male X chromosome is silenced in the germline, then one expectation is that it should have a chromatin conformation consistent with decreased transcriptional activity. Chromatin structure can be regulated via differential modification of nucleosomal histone N termini ‘tails’, which includes acetylation, methylation, phosphorylation and ubiquitination (Strahl and Allis, 2000 Turner, 2000). Combinations of these modifications are proposed to comprise a ‘histone code’ that determines regional structural properties of chromatin (Strahl and Allis, 2000). The acetylation of lysine residues in the N termini of histones H3 and H4, as well as methylation of lysine 4 in histone H3, generally correlate with a transcriptionally active state. By contrast, methylation of lysine 9 in histone H3 correlates with transcriptional silencing and constitutive heterochromatin formation, and is required for binding of the heterochromatin protein HP1 (Strahl and Allis, 2000 Jenuwein, 2001). Some proteins containing a SET domain are histone methyltransferases that can methylate lysine 9 in histone H3 (Jenuwein, 2001 Jenuwein and Allis, 2001). The general scheme of a ‘histone code’ has probably undergone specific adaptations in different organisms, but overall remains strongly conserved.
We have used probes specific for histone modifications to study the chromatin organization of the X chromosome in germ cells of C. elegans males and hermaphrodites. Both germline-silenced and germline-expressing transgene arrays were used to monitor how the histone modification patterns on these large, extrachromosomal arrays correlate with expression competence. The spectrum of histone modifications on transgene arrays illustrate a consistent correlation with the expression competence of the array in early meiotic germ cells. Moreover, we present evidence that, relative to autosomes, the histones on the X chromosome in male germ cells show a marked reduction in modifications that correlate with transcriptional activation and are enriched in a modification that is associated with heterochromatin.
Strikingly, the X chromosomes in oogenic hermaphrodite germ cells also appear silenced in early meiotic prophase as assessed by their histone modification pattern. Oocyte-enriched genes on the X chromosomes are, on average, expressed at levels significantly lower than oocyte-enriched genes on autosomes. Transcription of several X-linked oocyte genes was only detected in very late meiotic prophase I in the female germline of hermaphrodites.
We also demonstrate that three types of unpaired autosomal sequences are competent to express genes and display activating chromatin modifications: extrachromosomal transgene arrays containing interspersed genomic DNA, unpaired autosomal duplications and the autosomal portions of X:autosome translocations. Each has histone modification patterns more similar to autosomes than to the X chromosome throughout meiosis. These results show that pairing is probably not required for gene expression during meiosis, and suggest that chromatin on autosomes may be refractory to germline silencing. We also show that silencing of the X chromosome is a conserved feature in nematode species with divergent modes of reproduction, and thus does not appear to be a consequence of hermaphroditism.
What's the difference between a free chromosome fragment and an extrachromosomal array? - Biology
By the end of this section, you will be able to do the following:
- List the different steps in prokaryotic transcription
- Discuss the role of promoters in prokaryotic transcription
- Describe how and when transcription is terminated
The prokaryotes, which include Bacteria and Archaea, are mostly single-celled organisms that, by definition, lack membrane-bound nuclei and other organelles. A bacterial chromosome is a closed circle that, unlike eukaryotic chromosomes, is not organized around histone proteins. The central region of the cell in which prokaryotic DNA resides is called the nucleoid region. In addition, prokaryotes often have abundant plasmids, which are shorter, circular DNA molecules that may only contain one or a few genes. Plasmids can be transferred independently of the bacterial chromosome during cell division and often carry traits such as those involved with antibiotic resistance.
Transcription in prokaryotes (and in eukaryotes) requires the DNA double helix to partially unwind in the region of mRNA synthesis. The region of unwinding is called a transcription bubble. Transcription always proceeds from the same DNA strand for each gene, which is called the template strand. The mRNA product is complementary to the template strand and is almost identical to the other DNA strand, called the nontemplate strand, or the coding strand. The only nucleotide difference is that in mRNA, all of the T nucleotides are replaced with U nucleotides ((Figure)). In an RNA double helix, A can bind U via two hydrogen bonds, just as in A–T pairing in a DNA double helix.
Figure 1. Messenger RNA is a copy of protein-coding information in the coding strand of DNA, with the substitution of U in the RNA for T in the coding sequence. However, new RNA nucleotides base pair with the nucleotides of the template strand. RNA is synthesized in its 5′-3′ direction, using the enzyme RNA polymerase. As the template is read, the DNA unwinds ahead of the polymerase and then rewinds behind it.
The nucleotide pair in the DNA double helix that corresponds to the site from which the first 5′ mRNA nucleotide is transcribed is called the +1 site, or the initiation site. Nucleotides preceding the initiation site are denoted with a “-” and are designated upstream nucleotides. Conversely, nucleotides following the initiation site are denoted with “+” numbering and are called downstream nucleotides.
Initiation of Transcription in Prokaryotes
Prokaryotes do not have membrane-enclosed nuclei. Therefore, the processes of transcription, translation, and mRNA degradation can all occur simultaneously. The intracellular level of a bacterial protein can quickly be amplified by multiple transcription and translation events that occur concurrently on the same DNA template. Prokaryotic genomes are very compact, and prokaryotic transcripts often cover more than one gene or cistron (a coding sequence for a single protein). Polycistronic mRNAs are then translated to produce more than one kind of protein.
Our discussion here will exemplify transcription by describing this process in Escherichia coli, a well-studied eubacterial species. Although some differences exist between transcription in E. coli and transcription in archaea, an understanding of E. coli transcription can be applied to virtually all bacterial species.
Prokaryotic RNA Polymerase
Prokaryotes use the same RNA polymerase to transcribe all of their genes. In E. coli, the polymerase is composed of five polypeptide subunits, two of which are identical. Four of these subunits, denoted α, α, β, and β‘, comprise the polymerase core enzyme. These subunits assemble every time a gene is transcribed, and they disassemble once transcription is complete. Each subunit has a unique role the two α-subunits are necessary to assemble the polymerase on the DNA the β-subunit binds to the ribonucleoside triphosphate that will become part of the nascent mRNA molecule and the β‘ subunit binds the DNA template strand. The fifth subunit, σ, is involved only in transcription initiation. It confers transcriptional specificity such that the polymerase begins to synthesize mRNA from an appropriate initiation site. Without σ, the core enzyme would transcribe from random sites and would produce mRNA molecules that specified protein gibberish. The polymerase comprised of all five subunits is called the holoenzyme.
A promoter is a DNA sequence onto which the transcription machinery, including RNA polymerase, binds and initiates transcription. In most cases, promoters exist upstream of the genes they regulate. The specific sequence of a promoter is very important because it determines whether the corresponding gene is transcribed all the time, some of the time, or infrequently. Although promoters vary among prokaryotic genomes, a few elements are evolutionarily conserved in many species. At the -10 and -35 regions upstream of the initiation site, there are two promoter consensus sequences, or regions that are similar across all promoters and across various bacterial species ((Figure)). The -10 sequence, called the -10 region, has the consensus sequence TATAAT. The -35 sequence has the consensus sequence TTGACA. These consensus sequences are recognized and bound by σ. Once this interaction is made, the subunits of the core enzyme bind to the site. The A–T-rich -10 region facilitates unwinding of the DNA template, and several phosphodiester bonds are made. The transcription initiation phase ends with the production of abortive transcripts, which are polymers of approximately 10 nucleotides that are made and released.
Figure 2. The σ subunit of prokaryotic RNA polymerase recognizes consensus sequences found in the promoter region upstream of the transcription start site. The σ subunit dissociates from the polymerase after transcription has been initiated.
Link to Learning
View this MolecularMovies animation to see the first part of transcription and the base sequence repetition of the TATA box.
Elongation and Termination in Prokaryotes
The transcription elongation phase begins with the release of the σ subunit from the polymerase. The dissociation of σ allows the core enzyme to proceed along the DNA template, synthesizing mRNA in the 5′ to 3′ direction at a rate of approximately 40 nucleotides per second. As elongation proceeds, the DNA is continuously unwound ahead of the core enzyme and rewound behind it. The base pairing between DNA and RNA is not stable enough to maintain the stability of the mRNA synthesis components. Instead, the RNA polymerase acts as a stable linker between the DNA template and the nascent RNA strands to ensure that elongation is not interrupted prematurely.
Prokaryotic Termination Signals
Once a gene is transcribed, the prokaryotic polymerase needs to be instructed to dissociate from the DNA template and liberate the newly made mRNA. Depending on the gene being transcribed, there are two kinds of termination signals. One is protein-based and the other is RNA-based. Rho-dependent termination is controlled by the rho protein, which tracks along behind the polymerase on the growing mRNA chain. Near the end of the gene, the polymerase encounters a run of G nucleotides on the DNA template and it stalls. As a result, the rho protein collides with the polymerase. The interaction with rho releases the mRNA from the transcription bubble.
Rho-independent termination is controlled by specific sequences in the DNA template strand. As the polymerase nears the end of the gene being transcribed, it encounters a region rich in C–G nucleotides. The mRNA folds back on itself, and the complementary C–G nucleotides bind together. The result is a stable hairpin that causes the polymerase to stall as soon as it begins to transcribe a region rich in A–T nucleotides. The complementary U–A region of the mRNA transcript forms only a weak interaction with the template DNA. This, coupled with the stalled polymerase, induces enough instability for the core enzyme to break away and liberate the new mRNA transcript.
Upon termination, the process of transcription is complete. By the time termination occurs, the prokaryotic transcript would already have been used to begin synthesis of numerous copies of the encoded protein because these processes can occur concurrently. The unification of transcription, translation, and even mRNA degradation is possible because all of these processes occur in the same 5′ to 3′ direction, and because there is no membranous compartmentalization in the prokaryotic cell ((Figure)). In contrast, the presence of a nucleus in eukaryotic cells precludes simultaneous transcription and translation.
Figure 3. Multiple polymerases can transcribe a single bacterial gene while numerous ribosomes concurrently translate the mRNA transcripts into polypeptides. In this way, a specific protein can rapidly reach a high concentration in the bacterial cell.
Link to Learning
Visit this BioStudio animation to see the process of prokaryotic transcription.
In prokaryotes, mRNA synthesis is initiated at a promoter sequence on the DNA template comprising two consensus sequences that recruit RNA polymerase. The prokaryotic polymerase consists of a core enzyme of four protein subunits and a σ protein that assists only with initiation. Elongation synthesizes mRNA in the 5′ to 3′ direction at a rate of 40 nucleotides per second. Termination liberates the mRNA and occurs either by rho protein interaction or by the formation of an mRNA hairpin.
Which subunit of the E. coli polymerase confers specificity to transcription?
Repression of HR
Recently, it has become clear that telomeres also need to be protected from inappropriate homologous recombination. There are three types of HR that have detrimental outcomes at chromosome ends. The first is homologous recombination between sister telomeres, referred to as Telomere Sister Chromatid Exchange (T-SCE). T-SCEs could be detrimental to cells if the exchanged sequences are not equal in length. One sister telomere could become lengthened at the expense of the other. The exchange between sister telomeres can be detected by chromosome-orientation fluorescent in situ hybridization (CO-FISH) (for review, see Bailey et al. 2004). In the CO-FISH method, newly synthesized DNA is degraded and the remaining parental strands are detected with single-stranded probes. At telomeres, one parental strand is composed of 5′-TTAGGG-3′ repeats and the other has only 5′-CCCTAA-3′ sequences, allowing the two strands to be distinguished in metaphase chromosomes. After semiconservative replication, each chromatid in a metaphase chromosome will have a G-strand signal at one end and a C-strand signal at the other. A chromatid end displaying both C-strand and G-strand signals indicates that a T-SCE event has occurred at this end. In normal mouse and human cells, T-SCE is not frequent, but ALT cells, which maintain their telomeres by a recombination pathway, have very frequent T-SCE (Bailey et al. 2004 Bechter et al. 2004 Londono-Vallejo et al. 2004). The difference may be that T-SCEs are normally repressed and that ALT cells have loosened their control of HR at telomeres, allowing them to maintain their telomeres and therefore to survive (for review, see Neumann and Reddel 2002).
A second HR reaction that threatens telomeres is referred to as t-loop HR (Wang et al. 2004) (Fig. 5). T-loops are at risk for resolution by Holliday junction (HJ) resolvases because an HJ could be formed if the 5′ end of the telomere base pairs with the displacement loop (D loop). Branch migration in the direction of the centromere could generate a double HJ and resolution of this structure with crossover would delete the whole loop segment, leaving a drastically shortened telomere at the chromosome end. T-loop HR was discovered through a separation of function mutant of TRF2, TRF2 ΔB , which protects telomeres from NHEJ but induces sudden telomere truncations. These deletions are dependent on two proteins implicated in HR, the Mre11 recombination repair complex and XRCC3, a Rad51 paralog associated with HJ resolution activity. Cells expressing TRF2 ΔB also contain extrachromosomal telomeric DNA that is circular. On two-dimensional gels, these circles show a broad size distribution consistent with their representing the loop part of the t-loops. How the N terminus of TRF2 represses t-loop HR has not been established. As unperturbed cells contain small amounts of circular telomeric DNA, suppression of the t-loop HR reaction at telomeres may be incomplete. The control of t-loop HR appears to be further relaxed in ALT cells, which contain abundant telomeric circles (Cesare and Griffith 2004 Wang et al. 2004). As T-SCE and t-loop HR are similar reactions (one taking place in cis, the other in trans), their prevalence in ALT cells may be due to loss of a repressor that controls both.
There may be a third type of HR with detrimental outcomes—the recombination between a telomere and interstitial telomeric DNA. Chromosome internal telomeric DNA is not frequent in human cells, but in many other vertebrates, such sequences are abundant throughout the chromosomes. Recombination between telomeres and these elements could generate terminal deletions, extrachromosomal fragments, inversions, and translocations. This type of recombination appears to take place in mouse cells lacking ERCC1, which generate large extrachromosomal elements that contain a single stretch of telomeric DNA, presumably at a chromosome internal site (Zhu et al. 2003). These elements, referred to as Telomeric DNA-containing Double Minute chromosomes (TDMs) could be formed by recombination between a telomere and interstitial telomeric DNA on the same chromosome. Perhaps shelterin carries ERCC1/XPF is on its “tool-belt” to prevent inappropriate recombination events.
Present address: The Institute for Genomic Research, 9712 Medical Center Drive, Rockville, Maryland, 20850, USA
L. Eichinger, J. A. Pachebat, G. Glöckner, M.-A. Rajandream and R. Sucgang: *These authors contributed equally to this work
Center for Biochemistry and Center for Molecular Medicine Cologne, University of Cologne, Joseph-Stelzmann-Str. 52, 50931, Cologne, Germany
L. Eichinger, J. A. Pachebat, B. Tunggal, F. Rivero, P. Farbrother & A. A. Noegel
Laboratory of Molecular Biology, MRC Centre, CB2 2QH, Cambridge, UK
J. A. Pachebat, S. Kummerfeld, M. Madera, B. A. Konfortov, A. T. Bankier, M. Madan Babu, A. Wardroper, R. R. Kay & P. H. Dear
Genome Analysis, Institute for Molecular Biotechnology, Beutenbergstr. 11, D-07745, Jena, Germany
G. Glöckner, K. Szafranski, R. Lehmann, M. Felder, A. Rosenthal & M. Platzer
The Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridgeshire, CB10 1SA, UK
M.-A. Rajandream, M. Berriman, N. Hamlin, R. Davies, D. Saunders, P. Davis, A. Kerhornou, N. Hall, N. Bason, C. Churcher, J. Cooper, A. Cronin, I. Goodhead, T. Mourier, A. Pain, D. Harper, H. Hauser, K. James, D. Johnson, A. Knights, K. Mungall, K. Oliver, C. Price, M. A. Quail, E. Rabbinowitsch, M. Sanders, S. Sharp, M. Simmonds, S. Spiegler, A. Tivey, B. White, D. Walker, J. Woodward & B. Barrell
Verna and Marrs McLean Department of Biochemistry and Molecular Biology, Baylor College of Medicine, Houston, Texas, 77030, USA
R. Sucgang, J. Song, G. Chen, X. Nie, L. Hemphill, B. Desany, M. Lu, R. Lindsay, J. Ma & A. Kuspa
Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas, 77030, USA
Q. Xu, E. Sodergren, N. van Driessche, G. Shaulsky, G. Weinstock, R. Gibbs & A. Kuspa
Graduate Program in Structural and Computational Biology and Molecular Biophysics, Baylor College of Medicine, Houston, Texas, 77030, USA
Human Genome Sequencing Center, Baylor College of Medicine, Houston, Texas, 77030, USA
E. Sodergren, D. Muzny, M. Quiles, H. Loulseged, J. Hernandez, D. Steffen, G. Weinstock & R. Gibbs
Section of Cell and Developmental Biology, Division of Biology, University of California, La Jolla, California, 92093, San Diego, USA
R. Olsen, C. Anjard & W. F. Loomis
dictyBase, Center for Genetic Medicine, Northwestern University, 303 E Chicago Ave, Chicago, Illinois, 60611, USA
P. Gaudet, P. Fey, K. Pilcher, E. Just & R. L. Chisholm
Graduate School of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Ibaraki, 305-8572, Japan
T. Morio, H. Urushihara & Y. Tanaka
Adolf-Butenandt-Institute/Cell Biology, Ludwig-Maximilians-University, 80336, Munich, Germany
Biochemistry Department, University of Cambridge, Cambridge, CB2 1QW, UK
Division of Biological Sciences, Graduate School of Science, Hokkaido University, 060-0810, Sapporo, Japan
Unité de Genomique des Microorganismes Pathogenes, Institut Pasteur, 28 rue du Dr Roux, 75724, Cedex 15, Paris, France
Department of Biology, University of York, York, YO10 5YW, UK
MRC Cancer Cell Unit, Hutchison/MRC Research Centre, Hills Road, CB2 2XZ, Cambridge, UK
Centre for Genetic Resource Information, National Institute of Genetics, Mishima, Shizuoka, 411-8540, Japan
Department of Medical Genome Sciences, Graduate School of Frontier Sciences, The University of Tokyo, Minato, Tokyo, 108-8639, Japan
Institut für Pharmazeutische Biologie, Universität Frankfurt (Biozentrum), 60439, Frankfurt am Main, Germany
Department of Molecular Biology, Princeton University, Princeton, New Jersey, 08544-1003, USA
School of Life Sciences, University of Dundee, Dow Street, Dundee, DD1 5EH, UK