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Telomeres and daughter strands

Telomeres and daughter strands


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How is the primer present in the daughter, leading strand replaced? I've circled the primer in question.

I don't see how this primer can be replaced by DNA polymerase; there is no free 3' end for DNA pol to act on.

My book, however, says there is no telomere problem with the leading strand.

Other sources say there is a telomere problem.

So what is it?


Replication doesn't start at the very end of a chromosome, so there is no problem with leading strand synthesis. It's probably easiest to see if the other half of the image was there:

Forgive the lion; plain white paper seems to be in short supply. Also my writing. Too much tea this morning. Hopefully you can see that leading strand synthesis can continue right to of the end of the chromosome as the fork progresses. The leading primers at the replication origin can be replaced once the first Okazaki fragment from the other replication fork is synthesized.

There is, however, a problem, aptly termed the end replication problem, with lagging stand synthesis:

The dots just mean the chromosome continues in that direction. Important to notice is the lagging strand RNA primer at the end of the chromosome. When this is removed, there is apparently no way to synthesize DNA there. Repeated replication cycles progressively shorten the chromosome and cause the Hayflick limit to cell division (the number of times a cell can divide before dying). To solve this problem, some cell types express an enzyme called telomerase. This is an RNA directed DNA polymerase that adds deoxyribonucleotide repeats to the 3' end of a DNA strand, using its own internal RNA template, to prevent information loss.


I think the confusion is coming from the way this picture is drawn. The leading strand does not begin at the end of a chromosome, there is in fact more DNA to the right of your picture that is not shown. This picture is showing the left half of a replication bubble.


I think I understand this. Replication will occur bidirectionally from the the origin inside the replication bubble. If we focus just on 1 parent strand being replicated, on one side of the origin, the daughter stand replication will be continuous. However, on the other side of the same parent strand, the replication will require Okazaki fragments.

So for the same daughter strand, on one side of the replication origin, it will be leading, and on the other side of the origin it will be lagging!

Back to the telomere shortening (end replication problem), this will mean that each daughter strand will end up too short on it's 5' end after the RNA primer (Okazaki fragment) is removed.


Telomeres

Each eukaryotic chromosome consists of a single molecule of DNA associated with a variety of proteins.

The DNA molecules in eukaryotic chromosomes are linear i.e., have two ends. (This is in contrast to such bacterial chromosomes as that in E. coli that is a closed circle, i.e. has no ends.)

The DNA molecule of a typical chromosome contains

  • a linear array of genes (encoding proteins and RNAs) interspersed with
  • much noncoding DNA.

Included in the noncoding DNA are

  • long stretches that make up the centromere and
  • long stretches at the ends of the chromosome, the telomeres.

Telomeres are crucial to the life of the cell. They keep the ends of the various chromosomes in the cell from accidentally becoming attached to each other.

The telomeres of humans consist of as many as 2000 repeats of the sequence
5' GGTTAG 3'.


Defective telomeres are now being linked to dozens of diseases, including many types of cancer

A) Shelterin recruits accessory proteins to the telomeres that facilitate the complex process of telomere copying and maintenance associated with cell multiplication. B) Representative images of metaphase chromosomes of cells with functional telomeres (top) and dysfunctional telomeres that result in chromosomal fusions (bottom). Credit: CNIO

Studying telomeres, the structures that protect the ends of chromosomes, has become a key issue in biology. In recent years, not only has their relation to ageing been confirmed defective telomeres seem to be linked to more and more illnesses, including many types of cancer. The review published by Paula Martínez and María Blasco from the CNIO in Trends in Biochemical Sciences, stresses the importance of investigating these structures to improve diagnoses and develop possible treatments for many diseases. Telomeres, in the opinion of these researchers, will become increasingly important in clinical studies.

The chromosomes in every single cell are made up of DNA and shaped like strands, with a kind of protective cap at the end of each strand of DNA. Without this end protective cap, the DNA strands would chemically bond to other strands, i.e. the chromosomes would merge and that would be lethal for the cell. The structures that prevent this catastrophe are the telomeres. They were discovered in the 1930s but decades elapsed before someone decided to study them in any depth and since the late 1990s they have always been on the cutting edge of biology research. Biologists are often surprised by their amazing and unexpected complexity, and their health-related significance.

"The biology of telomeres is extremely complex and the more we discover the more we realise what remains to be discovered", says Paula Martínez from CNIO's Telomere and Telomerase Group. "What surprises me most is the high number of factors we are finding that are essential to the preservation of telomeres and, above all, the precise coordination that is required between them all".

The fact that telomeres have been tightly preserved throughout the evolutionary tree -in most eukaryotes: vertebrates, plants and even unicellular organisms such as yeast- indicates their importance. In addition to preventing the merger of chromosomes, telomeres are needed to prevent the loss of genetic information each time a cell divides.

Preventing Information Loss

When a cell replicates, the molecular machinery in charge of duplicating the chromosomes - so that each daughter cell has a copy -cannot reach the tip. This is inherently impossible due to the way the DNA replication machinery works, and it implies that any genetic material at the end of a chromosome with significant information for the cell would be lost. Telomeres prevent this from happening: they consist of a DNA sequence that does not contain genes and that is repeated numerous times- in humans and other species the sequence is TTAGGG the letters correspond to three of the building blocks that make up the DNA: thymine, adenine and guanine.

Consequently, the shortening of the DNA with every division is not significant. At least not until a certain limit is reached. When the telomeres become too short, we see the problems associated with ageing: cells reach a point where they interpret critically short telomeres as irreparable damage and react by no longer dividing, which prevents tissue from regenerating.

This happens in healthy cells but not in cancer cells. There is an enzyme, telomerase, which is capable of lengthening the telomeres de novo. This enzyme is not present in most cells of an adult organism but it is active in tumour cells. By repairing the telomeres, the telomerase enables cancer cells to proliferate and become virtually immortal.

This link to ageing and cancer, has led to the intense study of telomere-based strategies to combat cancer and diseases associated with ageing. Blasco's group has recently shown that it is possible to make cancer cells mortal by acting on the telomeres.

Zooming In To The Tip Of The Buffer

The above-mentioned description of telomeres however is a simplified version of the story. We now know that there is a protective structure enveloping telomeric DNA consisting of six proteins known as shelterins, which are crucial. Another more recent discovery is that there are proteins that, although not in the telomeres themselves, interact with them at specific times to enable them to perform their functions.

These proteins enable the telomeres to unwind, for example because, the sequence repeated in telomeres, TTAGGG, ends in a single strand of DNA that curves forming a loop and connects to the original strand of the double chain forming a triple chain. "Yes, it is very complicated", admits Martínez. "Structures of up to four chains of DNA can form".

When a cell divides, the telomeres are also replicated. This implies that the end loop must unwind first and then form again. This process also contributes to the shortening of telomeres and we now know that some of the shelterins as well as other associated proteins that interact with telomeres are key elements in this process.

According to Martínez, "there is now more evidence about relationship between telomere maintenance and several illnesses".

Telomere syndromes, or telomeropathies, have been identified in patients with mutations of the telomerase enzyme. This group includes, for example, pulmonary fibrosis and problems related to the malfunction of the bone marrow. A direct relationship between telomere dysfunctions and many types of cancer has also been found. More recently, we have also discovered that mutations of the proteins that protect telomeric DNA, the shelterins, and those that interact with the telomeres, are linked to various diseases, such as dyskeratosis congenita, Hoyeraal-Hreidarsson syndrome or Revesz syndrome.

"These discoveries underline the plethora of components and pathways that control telomere functions", write the authors in the paper. "In the future, research will bring to light more unknown factors that will improve our understanding of the mechanisms governing cancer and syndromes linked to the shortening of telomeres. We hope that this knowledge will be transferred to the clinic in order to improve the diagnosis and treatment of diseases".


RESULTS

Telomerase RNA (TERC) knockout cells engage the alternative lengthening of telomeres (ALT) pathway during crisis, a stage of massive cell death

To test whether the ALT pathway can be activated by telomerase inhibition in human telomerase positive transformed cells, we generated telomerase functional RNA (TERC) knockouts (KO) in the SW39 cell line using the CRISPR/Cas9 genome editing system (Figure 1A). SW39 cells are immortal telomerase positive cells derived from human lung fibroblasts (IMR90 cells) which were stably transfected with SV40 large-T antigen ( 26). SW39 cells were transformed with a Cas9/TERC targeting guide RNA. Single clones were screened by PCR and a droplet digital PCR-based telomere repeat amplification protocol (ddTRAP) ( 25) to verify the deletion of the TERC locus and the loss of telomerase activity. SW39 TERC KO cells proliferated ∼21–24 population doublings (PDs) before entering crisis, a period that is accompanied by a balance between cell growth and massive cell death ( Supplementary Figure S1A ). During crisis, cells acquire many genetic alterations in addition to the edited loss of the TERC gene. We predicted that only a small fraction of cells would escape from crisis by acquiring a telomerase-independent telomere maintenance mechanism. A modification of a fluctuation analysis using SW39 TERC KO cells resulted in emergence of three survival clones out of 12 million cells in crisis resulting in a frequency of immortalization = 2.5 × 10 −7 ( Supplementary Figure S1B ), confirming that overcoming the crisis event occurs at a low frequency.

Telomerase inhibition in telomerase-positive cells onsets the alternative lengthening of telomeres to overcome the crisis. (A) Overview of ALT cell line generation from telomerase positive transformed cells (SW39). SW39 cells were introduced with Cas9/TERC targeting guide RNA and single cell clones were isolated and analyzed. (B) Terminal Restriction Fragment (TRF) analysis in SW39 (parental cell) and SW39 TERC KO survival clones (post-crisis cells). Digested DNA was run on 0.85% agarose gels. M indicates for the marker. (C) Droplet digital PCR based telomere repeat amplification (ddTRAP) analysis in SW39, SW39 TERC KO survival clones (SW39 ALT1, ALT2, and ALT3), SW26 and SW13 cells (mean ± SD n = 3). (D) C-circle analysis in SW39 and SW39 ALT survival clones. One hundred nanograms of digested DNA from SW39 or SW39 ALT survival clones were used in the phi29 polymerase reaction.

Telomerase inhibition in telomerase-positive cells onsets the alternative lengthening of telomeres to overcome the crisis. (A) Overview of ALT cell line generation from telomerase positive transformed cells (SW39). SW39 cells were introduced with Cas9/TERC targeting guide RNA and single cell clones were isolated and analyzed. (B) Terminal Restriction Fragment (TRF) analysis in SW39 (parental cell) and SW39 TERC KO survival clones (post-crisis cells). Digested DNA was run on 0.85% agarose gels. M indicates for the marker. (C) Droplet digital PCR based telomere repeat amplification (ddTRAP) analysis in SW39, SW39 TERC KO survival clones (SW39 ALT1, ALT2, and ALT3), SW26 and SW13 cells (mean ± SD n = 3). (D) C-circle analysis in SW39 and SW39 ALT survival clones. One hundred nanograms of digested DNA from SW39 or SW39 ALT survival clones were used in the phi29 polymerase reaction.

To understand how these surviving clones escaped from crisis, we first measured telomere length using TRF analysis to check whether these clones escaped from crisis by elongating their telomere length. All three clones (herein after termed SW39 ALT1, 2 and 3) exhibited elongated and heterogeneous (long and short) telomere lengths compared to the parental cells (SW39) (Figure 1B). We performed ddTRAP analysis to assess telomerase activity in SW39 ALT cells, SW39, and its sister spontaneously occurring ALT cell lines, SW26 and SW13 as controls ( 26). As expected, SW39 showed robust telomerase activity, but no telomerase activity was detected in SW26 and SW13 (Figure 1C). In addition, all TERC KO surviving clones of SW39 had no detectable telomerase activity, suggesting that SW39 TERC KO cell lines acquired a telomerase-independent telomere maintenance mechanism.

C-rich circular extra-chromosomal DNAs (C-circles) and ALT-associated PML bodies (APBs) are accepted biomarkers of ALT cells ( 21). To determine if SW39 ALT cells have these ALT biomarkers, we first performed the C-circle assay in SW39 ALT cells and compared them to SW39 (Figure 1D). We detected a significant increase in C-circle levels in all SW39 ALT clones but not in the parental telomerase positive cell line SW39. Another biomarker of the ALT phenotype is the presence of ALT associated PML bodies (APBs). APB-positive cells were observed in SW39 ALT cells whereas the parental cells were APB-negative ( Supplementary Figure S1C ). In contrast, we did not detect any alteration of telomeric repeat-containing RNA (TERRA) levels in SW39 ALT cells ( Supplementary Figure S1D ), which has previously been suggested as another ALT pathway-associated factor ( 27). The phenotypes in SW39 ALT cells, including the heterogeneous and elongated telomere length, and presence of C-circle and APBs resemble those of ALT cells. These results indicate that telomerase inhibition in human telomerase positive transformed cells can result in engagement of ALT but at a low frequency.

ALT cells have excessively long overhangs

Telomeric overhangs are the binding site for telomerase and are elongated throughout S-phase in telomerase positive human cells ( 28). Moreover, telomere overhangs are thought to be important as the elongation site for ALT cells as well as telomerase positive cells ( 29). The exposed 3΄ telomeric end overhang is believed to invade into the sister chromatid, other chromosome ends or extra-chromosomal telomeric DNA repeats that can be used as templates to then elongate the ends via recombination mediated DNA replication ( 30). Therefore, we tested whether the alterations in telomere maintenance mechanism resulted in overhang length changes. Duplex-specific nuclease (DSN) digests double-stranded DNA into <10 bp fragments while leaving the single stranded telomeric overhangs intact (Figure 2A) ( 22, 31). Using the DSN method, we measured the overhang length in SW39 and SW39 ALT1 cells. Overhang length in SW39 ALT1 cells are variable from 40 nt to 400 nt, whereas parental SW39 cells have 65 nt to 140 nt overhang lengths consistent with previous reports (overhang length in telomerase positive cells: 60–150 nt) ( 22, 24, 28) (Figure 2B). These results show that ALT cells can have excessively long overhang populations, possibly due to its less well controlled elongation mechanism compared to telomerase-mediated elongation as previously speculated ( 29, 32, 33).

Excessively elongated lagging overhangs in ALT cells. (A) Illustration demonstrating Duplex-specific nuclease (DSN) overhang assay. DSN enzyme digests double stranded DNA and leaves the single stranded TTAGGG sequences present in the telomeric G-overhangs (3΄-overhangs). (B) DSN overhang assay in SW39 and SW39 ALT1 cells. Digested DNA (15 μg each) was run on 1.2% alkaline agarose gels. C-rich sequence probes were used for detecting G-overhangs. Right: Lines traces of SW39 (black) and SW39 ALT1 (red) gel lanes. (C) Illustration demonstrating CsCl separation of telomeric leading and lagging strands. Cells were usually incubated with IdU (5-Iodo-2΄-deoxyuridine) for 20 h (asynchronous condition) and then leading and lagging strands were separated using CsCl centrifugation. (D) CsCl separation of leading versus lagging strands in SW39 and SW39 ALT1 cells. a.u. represents arbitrary unit. (E) In-gel hybridization of telomeres in SW39 and SW39 ALT1 cells after leading/lagging strand separation. Native gels were hybridized with a C-rich probe to detect the amount of G-overhangs. Denatured gels were hybridized with a G-rich probe to measure the amount of total telomeric DNAs. Dashed lines indicate the quantified area. (F) Quantification of G-overhang intensity in E. Relative ratio of native signal to denatured signal was calculated (mean ± SEM n = 2) ** indicates P < 0.01 and n.s. indicates non-significant (unpaired Student's t test).

Excessively elongated lagging overhangs in ALT cells. (A) Illustration demonstrating Duplex-specific nuclease (DSN) overhang assay. DSN enzyme digests double stranded DNA and leaves the single stranded TTAGGG sequences present in the telomeric G-overhangs (3΄-overhangs). (B) DSN overhang assay in SW39 and SW39 ALT1 cells. Digested DNA (15 μg each) was run on 1.2% alkaline agarose gels. C-rich sequence probes were used for detecting G-overhangs. Right: Lines traces of SW39 (black) and SW39 ALT1 (red) gel lanes. (C) Illustration demonstrating CsCl separation of telomeric leading and lagging strands. Cells were usually incubated with IdU (5-Iodo-2΄-deoxyuridine) for 20 h (asynchronous condition) and then leading and lagging strands were separated using CsCl centrifugation. (D) CsCl separation of leading versus lagging strands in SW39 and SW39 ALT1 cells. a.u. represents arbitrary unit. (E) In-gel hybridization of telomeres in SW39 and SW39 ALT1 cells after leading/lagging strand separation. Native gels were hybridized with a C-rich probe to detect the amount of G-overhangs. Denatured gels were hybridized with a G-rich probe to measure the amount of total telomeric DNAs. Dashed lines indicate the quantified area. (F) Quantification of G-overhang intensity in E. Relative ratio of native signal to denatured signal was calculated (mean ± SEM n = 2) ** indicates P < 0.01 and n.s. indicates non-significant (unpaired Student's t test).

Lagging strands are preferentially extended in ALT cells

In telomerase positive cells, most leading and lagging strand overhangs are extended by telomerase after telomere replication, therefore their leading and lagging overhang lengths are similar ( 24). However, overhang lengths are totally different in telomerase negative cells [lagging strand overhang (∼100 nt in BJ normal fibroblast cells) > leading strand overhang (∼30 nt in BJ cells)]. Lagging overhangs are determined shortly after completion of DNA replication, whereas leading overhangs are processed throughout S/G2 in telomerase negative cells ( 23). We tested if these differences in leading versus lagging strands affected the ALT-mediated telomere extension process. Using cesium chloride (CsCl) centrifugation (Figure 2C and D), we separated leading and lagging strands in SW39 ALT1 cells and then measured the overhang signals using in-gel hybridization analysis (Figure 2E). We found that the overhang signal intensity ratio between leading versus lagging overhangs in SW39 ALT1 cell was 1:2.5 while the ratio in the SW39 cell line was 1:1. These results show that SW39 ALT1 cells have a different leading versus lagging strand overhang composition. One possible interpretation is that the lagging overhangs in SW39 ALT1 cells are significantly longer than leading overhangs, whereas SW39 cells have no differences in overhang lengths (Figure 2E). Collectively, these data support the idea that excessive telomere extensions in ALT cells occur more at lagging overhangs.

Telomerase reactivation reverses ALT-mediated telomere extension

We reintroduced the human TERC (TR) gene into SW39 ALT1 cells to test whether reacquiring telomerase activity abrogated the ALT phenotypes in SW39 ALT cells. For this, we introduced TR only (SW39 ALT1 + TR) or both TR and TERT (SW39 ALT1 + TT) into SW39 ALT1 cells. TR and TERT overexpressed SW39 (SW39 + TT) were used as a control. First, we determined that introduction of TR resulted in telomerase reactivation in SW39 ALT1 ( Supplementary Figure S2A ). Next, we compared the overhang signals. The overhang signals in SW39 ALT1 cells were much stronger compared to SW39+TT cells (Figure 3A and B). Thus, telomerase reactivation in SW39 ALT1 cells significantly reduced the overhang signals. The DSN method confirmed overhang length shortening by reactivation of telomerase. The excessively extended overhangs in SW39 ALT1 cells were reduced in cells reacquiring telomerase activity (Figure 3C). Moreover, the shortest telomere populations (<5 kb in TRF gels) in SW39 ALT1 cells were diminished upon telomerase reactivation ( Supplementary Figure S2B ). These results are consistent with the previous report that telomerase overexpression dramatically elongates the telomeres, rather than just maintaining the shorter telomere length ( 34). Lastly, we compared the overhang length after leading versus lagging separation. Telomerase reactivation (SW39 ALT1 + TT) completely diminished preferred lagging overhang elongation in SW39 ALT1 cells. The overhang intensity in leading and lagging strands in SW39 ALT1 + TT cells was almost identical to telomerase positive cells (Figure 3D and E). Collectively these results indicate that telomerase reactivation diminishes the ALT-mediated telomere extension processes.

Differential effects of telomerase reactivation on ATRX expression. (A) In-gel hybridization of telomeres in SW39 and SW39 ALT1 cells after introducing TERC or TERT gene expressing retrovirus. Cells were analyzed at 30 days post-infection. (B) Quantification of G-overhang intensity in A. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 3). (C) DSN overhang assay in SW39 ALT1, SW39 ALT1 expressing TERC and TERT, and SW39 cells. Digested DNA (10 μg each) was run on 1% alkaline agarose gels (M: marker). Right: Lines traces of SW39 (black), SW39 ALT1 (red) and SW39 ALT1 + TT (blue). (D) In-gel hybridization of leading and lagging strand telomeres in SW39 ALT1 and SW39 ALT1 expressing TERC and TERT cells. (E) Quantification of G-overhang intensity in D. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 3). (F) Western-blot for ATRX level from SW39, SW39 ALT1, SW39 ALT2, SW26 and SW13 cell lines. SW39 are telomerase positive and SW39 ALT1, SW39 ALT2, SW26 and SW13 are ALT cells. Lamin A/C was used as a loading control. (G) In-gel hybridization of telomeres in SW39 ALT1 after ATRX shRNA infection. Cells were analyzed 30 days post-infection. (H) Quantification of G-overhang intensity in G. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 3). (I) In-gel hybridization of leading and lagging strand telomeres in SW39 ALT1 cells after ATRX shRNA infection. Cells were analyzed 60 days post-infection. (J) Quantification of G-overhang intensity in I. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SD n = 3). (K) In-gel hybridization of leading and lagging strand telomeres in SW39 ALT2 and SW39 ALT2 expressing TERC and TERT cells. Cells were analyzed 30 days post-infection. (L) Quantification of G-overhang intensity in K. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 4). * indicates P < 0.05, ** indicates P < 0.01, *** indicates P < 0.001, **** indicates P < 0.0001 and n.s. indicates non-significant (unpaired Student's t test).

Differential effects of telomerase reactivation on ATRX expression. (A) In-gel hybridization of telomeres in SW39 and SW39 ALT1 cells after introducing TERC or TERT gene expressing retrovirus. Cells were analyzed at 30 days post-infection. (B) Quantification of G-overhang intensity in A. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 3). (C) DSN overhang assay in SW39 ALT1, SW39 ALT1 expressing TERC and TERT, and SW39 cells. Digested DNA (10 μg each) was run on 1% alkaline agarose gels (M: marker). Right: Lines traces of SW39 (black), SW39 ALT1 (red) and SW39 ALT1 + TT (blue). (D) In-gel hybridization of leading and lagging strand telomeres in SW39 ALT1 and SW39 ALT1 expressing TERC and TERT cells. (E) Quantification of G-overhang intensity in D. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 3). (F) Western-blot for ATRX level from SW39, SW39 ALT1, SW39 ALT2, SW26 and SW13 cell lines. SW39 are telomerase positive and SW39 ALT1, SW39 ALT2, SW26 and SW13 are ALT cells. Lamin A/C was used as a loading control. (G) In-gel hybridization of telomeres in SW39 ALT1 after ATRX shRNA infection. Cells were analyzed 30 days post-infection. (H) Quantification of G-overhang intensity in G. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 3). (I) In-gel hybridization of leading and lagging strand telomeres in SW39 ALT1 cells after ATRX shRNA infection. Cells were analyzed 60 days post-infection. (J) Quantification of G-overhang intensity in I. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SD n = 3). (K) In-gel hybridization of leading and lagging strand telomeres in SW39 ALT2 and SW39 ALT2 expressing TERC and TERT cells. Cells were analyzed 30 days post-infection. (L) Quantification of G-overhang intensity in K. Relative ratio of native signal to denatured signal was calculated (mean ± SD n = 4). * indicates P < 0.05, ** indicates P < 0.01, *** indicates P < 0.001, **** indicates P < 0.0001 and n.s. indicates non-significant (unpaired Student's t test).

Loss of ATRX facilitates lagging overhangs elongation

Loss of the ATRX gene and mutations in ATRX are hallmarks of ALT-immortalized cells ( 13– 15). To test whether the alteration in ATRX expression correlates with ALT onset in SW39 ALT cells, we first measured the level of ATRX in SW39 ALT cells and SW39 (Figure 3F and Supplementary Figure S3A ). As expected, SW39 cells expressed ATRX. The level of ATRX in SW39 ALT1 and SW39 ALT3 were similar to SW39, but ATRX levels in SW39 ALT2 were markedly reduced. We found that spontaneously derived ALT cell lines, SW26 and SW13 exhibited different ATRX expression levels SW26 was ATRX-negative, while SW13 was ATRX-positive (Figure 3F). These results support the idea that loss of ATRX expression is insufficient to promote ALT ( 15, 16). To directly test the effect of ATRX in ALT, we infected ATRX shRNA in ATRX-positive ALT cells (SW39 ALT1) and analyzed cells 60 days post-infection. We first performed the C-circle assay ( Supplementary Figures S3B and C ). Depletion of ATRX in SW39 ALT1 cells produced a 3-fold increase of C-circle levels indicating that ATRX depletion in ALT cells may promote ALT activity but is insufficient for engagement of ALT.

We next investigated the mechanism by which ATRX ablation promotes ALT phenotypes. We measured the overhang intensity in SW39 ALT1 after ATRX depletion. Overhang intensity was significantly (∼30%) increased in ATRX-depleted SW39 ALT1 cells (Figure 3G and H), whereas it was not altered in ATRX-depleted SW39 cells ( Supplementary Figure S3D and E ). Exonuclease I treatment completely eliminated the overhang signal in ATRX-depleted SW39 ALT1 (Figure 3G Exo1 + lanes). This suggests that the distinctly enhanced overhang signal does not originate from an increase of exposed internal single stranded telomeric DNA induced by ATRX depletion. We tested whether ATRX status specifically affected leading or lagging strand extension (Figure 3I and J). We measured the overhang intensity of leading and lagging strand in control and ATRX-depleted SW39 ALT1 cells. The lagging overhangs in SW39 ALT1 cells were further increased after ATRX-depletion despite having similar amounts of leading overhangs. Moreover, the effect of telomerase reactivation in ATRX-negative SW39 ALT2 cells was less dramatic compared to those in ATRX-positive SW39 ALT1 cells (Figure 3K and L). Collectively these data implicate that the ALT pathway can be mediated by preferential elongation of lagging overhangs that is partially suppressed by ATRX.

TERC KO in telomerase-positive cancer cells results in engagement of the ALT pathway

Next, we generated TERC KO in telomerase positive cancer cell lines to test whether the ALT pathway can be activated by telomerase inhibition in a variety of telomerase positive cancer cells. H1299 is a telomerase positive cancer cell line derived from a non-small cell lung cancer patient and after TERC KO H1299 cells divided ∼40 PDs before entering crisis (Figure 4A). We only obtained a single survival clone out of more than one hundred million cells in crisis resulting in a very low frequency of immortalization = 0.9 × 10 −8 ( Supplementary Figure S1B ). Using another cancer cell line HT1080 (fibrosarcoma mesenchymal origin) we could not obtain any ALT survival clones even though 100 million cells were maintained in crisis ( Supplementary Figure S4 ). Since HT1080 cells have intact p53, HT1080 TERC KO cells entered into a senescence-like stage in late PDs which is one explanation of why we did not obtain any long-term survival clones ( Supplementary Figure S4B ). Using the HAP1 (derived from chronic myelogenous leukemia) cell line again no ALT survival clones were obtained. Survival clone (H1299 ALT) had no detectable telomerase activity but possessed C-circles (Figure 4B), suggesting that H1299 ALT cells acquired a telomerase-independent telomere maintenance mechanism. Indeed, H1299 ALT cells exhibited elongated and heterogeneous telomere lengths compared to parental cells (H1299 and H1299 TERC KO) (Figure 4C).

TERC KO in H1299 cells results in engaging ALT pathway. (A) Overview of ALT cell line generation from telomerase positive cancer cells (H1299). H1299 TERC KO cells were generated by introducing with Cas9/TERC targeting guide RNA. H1299 TERC KO cells divided 40 population doublings before entering crisis. Single survival clone (H1299 ALT) was acquired. (B) DdTRAP analysis (bottom) and C-circle assay (upper) in H1299, H1299 TERC KO and H1299 ALT cells. (C) TRF analysis in H1299, H1299 TERC KO (pre-crisis, PD = 37) and H1299 ALT (post-crisis) cells. Digested DNA was run on 0.8% agarose gels. M indicates for the marker. (D) CsCl separation of leading and lagging strands in H1299 and H1299 ALT cells. a.u. represents arbitrary unit. (E) In-gel hybridization of leading and lagging strand telomeres in H1299 and H1299 ALT cells. (F) Quantification of G-overhang intensity in E. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SEM n = 2) ** indicates P < 0.01 and n.s. indicates non-significant (unpaired Student's t test).

TERC KO in H1299 cells results in engaging ALT pathway. (A) Overview of ALT cell line generation from telomerase positive cancer cells (H1299). H1299 TERC KO cells were generated by introducing with Cas9/TERC targeting guide RNA. H1299 TERC KO cells divided 40 population doublings before entering crisis. Single survival clone (H1299 ALT) was acquired. (B) DdTRAP analysis (bottom) and C-circle assay (upper) in H1299, H1299 TERC KO and H1299 ALT cells. (C) TRF analysis in H1299, H1299 TERC KO (pre-crisis, PD = 37) and H1299 ALT (post-crisis) cells. Digested DNA was run on 0.8% agarose gels. M indicates for the marker. (D) CsCl separation of leading and lagging strands in H1299 and H1299 ALT cells. a.u. represents arbitrary unit. (E) In-gel hybridization of leading and lagging strand telomeres in H1299 and H1299 ALT cells. (F) Quantification of G-overhang intensity in E. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SEM n = 2) ** indicates P < 0.01 and n.s. indicates non-significant (unpaired Student's t test).

We compared the overhang composition in H1299 and H1299 ALT cells after leading versus lagging separation (Figure 4D). Consistent with SW39 ALT cells, H1299 ALT cells possess longer lagging overhangs, whereas H1299 telomerase positive cells possess similar leading and lagging overhang composition (Figure 4E and F). Collectively, these results indicate that telomerase inhibition in human telomerase positive cancer cells can also result in engagement of ALT but at a very low frequency.

ALT cancer cells exhibit preferential elongation of lagging overhangs

We tested whether the preferential elongation of lagging overhangs in SW39 ALT and H1299 ALT cells is also present in established ALT cancer cells. We analyzed the U2OS cell line, an ALT cancer cell line derived from an osteosarcoma patient. U2OS cells contain canonical telomeric repeat sequences (86.5% of telomeres sequences are TTAGGG), whereas telomere sequences in other ALT cells consist of variant repeats ( 35). This enabled us to separate leading and lagging strand using CsCl centrifugation in U2OS cells (Figure 5A). CsCl separation enabled us to collect leading and lagging strands that have been fully replicated ( Supplementary Figure S5 ). The lagging overhang signal intensity in U2OS cells was longer than leading overhangs similar to SW39 ALT cells. To check the overhang ratio in cells without a telomere maintenance mechanism, we generated TERC KO cells derived from HeLa LT (with long telomeres in pre-crisis but without a telomere maintenance mechanism Figure 5B). Interestingly, we found that the overhang ratio in HeLa LT TERC KO cells was 1:2.8 similar to normal BJ cells ( 23) (Figure 5C and D). We next measured the C-overhangs in this panel of cells. C-overhangs may be due to telomere trimming, a telomere-loop excision process resulting from over-elongated telomere length or telomere uncapping problems ( 36– 38). Telomere trimming has been observed in ALT cells possibly induced by excessive telomere elongation ( 30, 39). We found that C-overhangs in U2OS cells are more abundant in lagging strands, whereas HeLa LT cells have a similar ratio between leading and lagging strands (Figure 5E and F). Interestingly, HeLa LT TERC KO cells have excessive amounts of C-overhangs in lagging strands, possibly due to longer G-overhangs in lagging strands. These data can be interpreted to suggest that telomere trimming processes in ALT cells more frequently occur in lagging strands compared to leading strands, whereas in telomerase positive cells telomere trimming events occur similarly in leading and lagging strands.

Lagging strand overhangs are preferentially elongated in U2OS and ASF1-depleted HeLa cells. (A) CsCl separation of leading and lagging strands in U2OS cells. a.u. represents arbitrary unit. (B) DdTRAP analysis (bottom) and C-circle assay (upper) in U2OS, HeLa LT and HeLa LT TERC KO cells. (C) In-gel hybridization of leading and lagging strand G-overhangs in U2OS, HeLa LT and HeLa LT TERC KO cells. (D) Quantification of G-overhang intensity in C. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SD n = 3). (E) In-gel hybridization of leading and lagging strand C-overhangs in U2OS, HeLa LT and HeLa LT TERC KO cells. (F) Quantification of C-overhang intensity in E. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SEM n = 2). (G) In-gel hybridization of leading and lagging strand telomeres in U2OS and U2OS expressing TERC and TERT cells. (H) Quantification of G-overhang intensity in G. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SD n = 4). (I) C-circle assay in U2OS and HeLa LT TERC KO cells after control or ASF1 siRNA (siCtrl, siASF1) transfection. Cells were analyzed 3 days post-transfection. (J) DSN overhang assay in HeLa LT TERC KO cells after control or ASF1 siRNA transfection. Right: Lines traces of control siRNA (black) and ASF1 siRNA (red). (K) In-gel hybridization of leading and lagging strand telomeres in HeLa LT TERC KO cells after control or ASF1 siRNA transfection. (L) Quantification of G-overhang intensity in K. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SEM n = 2). * indicates P < 0.05, ** indicates P < 0.01 and *** indicates P < 0.001 (unpaired Student's t test).

Lagging strand overhangs are preferentially elongated in U2OS and ASF1-depleted HeLa cells. (A) CsCl separation of leading and lagging strands in U2OS cells. a.u. represents arbitrary unit. (B) DdTRAP analysis (bottom) and C-circle assay (upper) in U2OS, HeLa LT and HeLa LT TERC KO cells. (C) In-gel hybridization of leading and lagging strand G-overhangs in U2OS, HeLa LT and HeLa LT TERC KO cells. (D) Quantification of G-overhang intensity in C. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SD n = 3). (E) In-gel hybridization of leading and lagging strand C-overhangs in U2OS, HeLa LT and HeLa LT TERC KO cells. (F) Quantification of C-overhang intensity in E. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SEM n = 2). (G) In-gel hybridization of leading and lagging strand telomeres in U2OS and U2OS expressing TERC and TERT cells. (H) Quantification of G-overhang intensity in G. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SD n = 4). (I) C-circle assay in U2OS and HeLa LT TERC KO cells after control or ASF1 siRNA (siCtrl, siASF1) transfection. Cells were analyzed 3 days post-transfection. (J) DSN overhang assay in HeLa LT TERC KO cells after control or ASF1 siRNA transfection. Right: Lines traces of control siRNA (black) and ASF1 siRNA (red). (K) In-gel hybridization of leading and lagging strand telomeres in HeLa LT TERC KO cells after control or ASF1 siRNA transfection. (L) Quantification of G-overhang intensity in K. Relative ratio of native signal to denatured signal in leading or lagging strands was calculated (mean ± SEM n = 2). * indicates P < 0.05, ** indicates P < 0.01 and *** indicates P < 0.001 (unpaired Student's t test).

We found that telomerase reactivation significantly decreased the lagging overhang in SW39 ALT1, whereas lagging overhangs were partially reduced in SW39 ALT2 (Figure 3D, E, K and L). This is similar to IMRB+TT cells reported previously ( 40). We further showed that the effect of telomerase reactivation was related to ATRX status. Consistent with SW39 ALT2 cells, the lagging overhangs were partially reduced when the telomerase (TERT and TERC) genes were ectopically introduced back into ATRX-negative U2OS cells (Figure 5G and H) ( 15).

A recent study showed that depletion of the histone chaperone ASF1 induced ALT phenotypes mediated by replication stress and the ATR-dependent checkpoint machinery ( 32). This led us to test whether the ASF1 depletion-induced ALT pathway was mediated by preferential elongation of lagging overhangs. HeLa LT TERC KO cells were transfected with siRNAs targeting ASF1a and ASF1b genes. Consistent with the previous study, ASF1 depletion moderately increased C-circle levels in HeLa LT TERC KO cells (Figure 5I). The DSN method revealed that the overhang lengths were also increased in ASF1-depleted HeLa LT TERC KO cells (Figure 5J). Moreover, we found that ASF1 depletion elongated lagging overhangs more than leading overhangs in HeLa LT TERC KO cells (Figure 5K and L), consistent with the phenotype observed in ALT cells.

C-circles are more abundant in lagging strand telomeres compared to leading strand telomeres

The excessive amounts of C-circles that exist in ALT cells may be important in ALT activity ( 15, 21, 32, 41, 42). To test whether the presence of C-circles has strand preference, we measured C-circle levels after CsCl separation. We performed a phi29 polymerase reaction for each fraction after CsCl centrifugation in U2OS cells (Figure 6A and B). C-circle levels peaked in leading and lagging fractions and mostly overlapped with the telomere DNA peaks as a singlet ((i) and (ii) in Figure 6C). When C-circle levels were quantified in leading strand versus lagging strand fractions, we found that C-circle levels from lagging strand fractions were more abundant compared to the leading strand fractions (leading:lagging = 1:1.63).

Lagging strand telomeres have more C-circle levels compare to leading strand telomeres. (A) Slot-blot image for CsCl separation. U2OS cells were incubated with IdU 20 h, then separated leading and lagging strand using CsCl centrifugation. Each fraction was dialyzed, and loaded in slot-blots to represent the amount of telomeric DNA. (B) Slot-blot image for C-circle assay for each fraction after CsCl separation. Each fraction was subjected to the phi29 polymerase reaction to measure C-circle levels. (C) Quantification of Telomeric DNA-A and C-circle assay-B. Black Diamond: telomeric DNA, Red circle: C-circle assay. a.u. represents arbitrary unit. (D) Predicted model for the strand-dependent existence of C-circles. (i) C-circle level in the leading strand fraction, (ii) C-circle level in the lagging strand fraction, (iii) C-circle level in the strands after crossover between sister-chromatids. Dashed lines indicate nascent or daughter strands, Solid lines indicate parental strands, Blue lines indicate G-rich sequences and Red lines indicate C-rich sequences.

Lagging strand telomeres have more C-circle levels compare to leading strand telomeres. (A) Slot-blot image for CsCl separation. U2OS cells were incubated with IdU 20 h, then separated leading and lagging strand using CsCl centrifugation. Each fraction was dialyzed, and loaded in slot-blots to represent the amount of telomeric DNA. (B) Slot-blot image for C-circle assay for each fraction after CsCl separation. Each fraction was subjected to the phi29 polymerase reaction to measure C-circle levels. (C) Quantification of Telomeric DNA-A and C-circle assay-B. Black Diamond: telomeric DNA, Red circle: C-circle assay. a.u. represents arbitrary unit. (D) Predicted model for the strand-dependent existence of C-circles. (i) C-circle level in the leading strand fraction, (ii) C-circle level in the lagging strand fraction, (iii) C-circle level in the strands after crossover between sister-chromatids. Dashed lines indicate nascent or daughter strands, Solid lines indicate parental strands, Blue lines indicate G-rich sequences and Red lines indicate C-rich sequences.

It has been proposed that C-circles mainly exist in the form of single stranded C-rich repeats with a sub-fraction existing as double stranded ( 21). If C-circles are a single stranded C-rich circular DNA without major G-rich sequences, it's density in IdU incorporated conditions should be two singlets ∼1.800 g/ml (CCC-IdU-AA) derived from lagging strand and ∼1.750 g/ml (CCCTAA) derived from leading strand and unreplicated DNA. However, we observed that C-circles peaks mostly overlapped with total telomere DNA peaks in leading, lagging and unreplicated fractions. One possible explanation is that C-circles may present in chromatid fractions possibly annealed with G-overhangs ((i) and (ii) in Figure 6D). The doublet peak between the leading and lagging fraction was also observed and may represent C-circles in crossover strands ((iii) in Figure 6C and D). These results show that C-circles are more abundant in lagging strands which may be used in the telomere elongation mechanism, such as via rolling circle amplification ( 21).

Telomere elongation in ALT cells mostly occurs during S phase

We next investigated the overhang length changes in a cell cycle dependent manner to identify which cell cycle phases are responsible for the ALT-mediated telomere elongation. Cells were synchronized at the G1/S phase by using double thymidine block, then released, and harvested at each cell cycle phase (S, G2 and G1 phase). The overhang intensity in U2OS cells peaked during S phase and gradually declined at the G2 phase (Figure 7A and B). The intensity of both leading and lagging overhangs peaked during S phase, but the increase of lagging overhang intensity was much higher than that of leading overhangs in U2OS cells (Figure 7A and B). These results demonstrate that ALT-mediated telomere elongation processes mostly occur during S phase and that lagging overhangs are preferentially and excessively elongated.

ALT pathways occur during S phase. (A) In-gel hybridization of leading and lagging strand telomeres in U2OS cells for each cell cycle phases. (B) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in A (mean ± SD n = 4). (C) In-gel hybridization of leading and lagging strand telomeres in U2OS cells for S phase with or without 30 μM of RAD51 inhibitor (RI-1) treatment. Cells were synchronized at G1/S phase by using double thymidine block and releases with or without RAD51 inhibitor, and then harvested at 6 hours post release. (D) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in C (mean ± SD n = 3). (E) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in HeLa LT cells ( Supplementary Figure S6A ) (mean ± SDs n = 3). (F) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in HeLa LT TERC KO cells ( Supplementary Figure S6B ) (mean ± SEM n = 2). (G) Schematic models for telomere elongation processes in telomerase positive and ALT cells * indicates P < 0.05, ** indicates P < 0.01, *** indicates P < 0.001, **** indicates P < 0.0001 and n.s. indicates non-significant (unpaired Student's t test).

ALT pathways occur during S phase. (A) In-gel hybridization of leading and lagging strand telomeres in U2OS cells for each cell cycle phases. (B) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in A (mean ± SD n = 4). (C) In-gel hybridization of leading and lagging strand telomeres in U2OS cells for S phase with or without 30 μM of RAD51 inhibitor (RI-1) treatment. Cells were synchronized at G1/S phase by using double thymidine block and releases with or without RAD51 inhibitor, and then harvested at 6 hours post release. (D) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in C (mean ± SD n = 3). (E) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in HeLa LT cells ( Supplementary Figure S6A ) (mean ± SDs n = 3). (F) Quantification of G-overhang intensity in relative ratio of native signal to denatured signal in HeLa LT TERC KO cells ( Supplementary Figure S6B ) (mean ± SEM n = 2). (G) Schematic models for telomere elongation processes in telomerase positive and ALT cells * indicates P < 0.05, ** indicates P < 0.01, *** indicates P < 0.001, **** indicates P < 0.0001 and n.s. indicates non-significant (unpaired Student's t test).

Previous studies showed that RAD51 (mitotic recombinase) depletion or inhibition results in reduction of G-overhangs ( 30) and interchromosomal telomeric recombination ( 43, 44) in ALT cells. To check whether RAD51-mediated recombination processes affect overhang generation during S phase, we treated U2OS cells with a small molecule RAD51 inhibitor ( 45). The RAD51 inhibitor treatment reduced the overhang signals in both leading and lagging strands during S phase (Figure 7C and D). These results demonstrate that recombination processes mediate the ALT pathway.

Next, we asked whether the preferential elongation in lagging strands is an ALT-specific phenotype. We compared these observations with telomerase positive cells by analyzing overhang intensity in HeLa LT cells at each cell cycle phase. Telomerase positive HeLa LT cells exhibited increased overhang intensity in both leading and lagging strands at S phase that gradually declined at G2 phase. The overhang intensity were increased up to 2-fold during S phase in both leading and lagging strands (Figure 7E and Supplementary Figure S6A ). These results are consistent with previous studies showing that telomerase extends telomere overhangs during S phase, and the CST-pol alpha complex fills in the C-strand during the G2 phase ( 28, 46). In contrast to ALT cells, the extension rate was similar for both leading and lagging strands in telomerase positive cells (Figure 7E and Supplementary Figure S6A ) ( 24). We also measured the cell cycle dependent overhang intensity changes in HeLa LT TERC KO cells to determine the overhang generation processes in cells without a telomere maintenance mechanism. HeLa LT TERC KO cells initially are both telomerase-negative and ALT-negative when derived from telomerase positive HeLa LT cells (Figure 5B). As shown in BJ fibroblasts ( 23), HeLa LT TERC KO cells exhibited the same lagging overhang intensity throughout the cell cycle, and that leading strand overhangs were processed during S phase (Figure 7F and Supplementary Figure S6B ). Collectively these results show that ALT telomeres are preferentially elongated at the lagging overhangs during S phase (Figure 7G).


Materials and methods

Cell culture and cell cycle synchronization

BJ foreskin fibroblasts were cultured at 37°C in 5% CO2 in high-glucose DMEM medium (Hyclone) containing 15% Cosmic calf serum (Hyclone). For synchronization, logarithmically growing cells were washed twice with 1× PBS, presynchronized with low serum (high-glucose DMEM with 0.1% fetal bovine serum, 20 mM HEPES) for 48 h, and then fed fresh medium containing 15% fetal bovine serum and 2 μg/mL aphidicolin (A.G. Scientific) for 24 h. Cells were then washed three times with 1× PBS, released into fresh medium for 2 h, and then labeled with 100 μM BrdU. For pulse-chase experiments, cells were washed twice with 1× PBS and chased with 100 μM thymidine.

HeLa cervical carcinoma cells were used for some experiments requiring short labeling times and a large number of cells. HeLa cells were cultured at 37°C in 5% CO2 in high-glucose DMEM medium with 10% Cosmic calf serum (Hyclone). Exponentially growing HeLa cells were synchronized with 2 mM thymidine (Sigma) for 19 h, washed three times with 1× PBS, and incubated with fresh medium for 9 h. Two millimolar thymidine was added again for 16 h. Cells were washed three times with 1× PBS and released into fresh medium with 100 μM BrdU for 0–10 h.

Genomic DNA isolation

Genomic DNA was purified by Qiagen Blood and Cell Culture Midi kit. Precipitated DNA was washed twice with 70% ethanol and suspended in 10 mM Tris-HCl (pH 8). DNA was dissolved overnight at 37°C.

Generation of telomere probes

Twenty-four-nucleotide probes containing six 32 P-dC or six 32 P-dG were synthesized as described (Herbert et al. 2003). A hypersensitive C probe was designed to incorporate 13 total radioactive nucleotides (nine 32 P-dC + four 32 P-dA). T3C3+3 (5′-TTTCCCTAA) was used instead of T3C3+9 (5′-TTTCCCTAACCCTAA) and annealed to GTU4 (5′-GGGUUAGGGUUAGGGUUAGGGAAA) for 1 min at 90°C, for 15 min at 20°C, and for 15 min at 17°C. The synthesis reaction (Herbert et al. 2003) was modified as follows: 3.1 μL of 8× Roche buffer M, 1 μL of annealed template oligo (1.7 pmol/μL), 1 μL of dTTP (1.25 mM stock, 50 μM final), 7 μL of 32 P-dCTP (3000 Ci/mmol), 4 μL of 32 P-dATP (3000 Ci/mmol), 7.9 μL of Millipore H2O, and 1 μL of Klenow (5 U/μL) were combined in a final volume of 25 μL. After extension for 30 min at 20°C and for 5 min at 95°C (to inactivate Klenow to prevent probe degradation upon uracil deglycosylase [UDG] treatment), the reaction was cooled to room temperature. UDG (0.5 μL 1 U/μL) was added to degrade the GTU template, incubated for 15 min at 37°C, and then inactivated for 10 min at 95°C. Free isotopes were then removed using an Illustra Microspin G-25 column (GE Healthcare).

CsCl separation of leading and lagging telomeric daughters

CsCl gradient separation was performed as described (Zhao et al. 2011b) with modifications. In brief, 500 μg of purified genomic DNA was digested with HinfI and RsaI overnight in 250 μL and terminated by the addition of EDTA to 10 mM. DNA was mixed with CsCl solution (density of 1.79 g/mL with 5 mM Tris-HCl at pH 8, 2 mM EDTA) to obtain a final density of ∼1.760–1.770 g/mL and added to a polyallomer quick-seal centrifuge tube (Beckman). Samples were centrifuged at 55,000 rpm for 20 h at 25°C using a VTi-80 vertical rotor (Beckman). Fractions of the sample were collected, and aliquots were denatured and hybridized with a telomere-specific probe on a slot blot to identify the fractions that contained telomere DNA. The corresponding densities were obtained by measuring the refractive index. Leading DNA was located at a density of 1.790–1.800 g/mL, lagging DNA was located at a density of 1.760–1.770 g/mL, and unreplicated DNA was located at a density of 1.740–1.750 g/mL. Four to five fractions of each peak were pooled and desalted by agarose dialysis (rocking on a plug of 2% agarose made with 10 mM Tris-HCl at pH 8 in a 50-mL tube for 1 h at room temperature) followed by ethanol precipitation. DNA was then dissolved in 10 mM Tris-HCl (pH 8).

Telomere overhang analysis by in-gel hybridization

Five micrograms of purified leading or lagging DNA was briefly run into an 0.8% agarose gel in 1× TAE so that telomeres were not significantly separated by size and remained as a relatively tight band. DNA that had been digested with 10 U of exonuclease I (20 U/μL) (Epicentre Biotechnologies) in a 20-μL reaction for 1 h at 37°C and terminated by adding 0.5 μL of EDTA (0.5 M stock) served as a negative control. The gel was dried, prehybridized (6× SSC, 5× Denhardt's solution, 0.5% [w/v] SDS), and hybridized with C probe for G overhangs under native conditions. Upon obtaining the signal from the native hybridization, the gel was denatured (0.5 M NaOH, 1.5 M NaCl for 1 h), rinsed three times with distilled water, neutralized (0.5 M Tris-HCl at pH 8, 1.5 M NaCl for 30 min), and then hybridized again with the C probe to obtain the denatured signal for total telomere input.

Overhang analysis by DSN assay

DSN assay was performed as described (Zhao et al. 2008, 2011a). In brief, 2–5 μg of purified leading versus lagging was digested with 2 U of DSN (Evrogen) for 2 h at 37°C. For negative controls, 10 U of ExoI (Epicentre Biotechnologies) was added to the genomic DNA for 1 h at 37°C prior to DSN digestion. DSN digestion was stopped by adding EDTA to a final concentration of 25 mM. For denaturing polyacrylamide gels, a 1:1 ratio of deionized formamide was added. DNA was then heated for 5 min at 65°C and loaded on a 6% polyacrylamide gel with 8 M urea. The gel was run at 15 V/cm in 0.5× TBE until the bromophenyl blue dye front-migrated two thirds into the length of the gel. DNA was electro-transferred onto a Amersham Hybond-N + membrane (GE Healthcare) with 0.5× TBE. The membrane was air-dried, UV-cross-linked, hybridized to a telomere-specific probe as described (Herbert et al. 2003), and exposed on a PhosphorImager (GE Healthcare). Alkaline agarose gels (1% agarose made with 50 mM NaOH, 1 mM EDTA) were run in 50 mM NaOH and 1 mM EDTA at 4°C with a low voltage (1–2 V/cm) until the dye migrated ∼6–8 cm. DNA was capillary-transferred to Amersham Hybond-XL (GE Healthcare) and processed as above. Average overhang sizes were calculated using the formula mean average length = ∑(Inti)/∑(Inti/MWi), where Inti is the signal intensity, and MWi is the molecular weight of the DNA at position i (Chai et al. 2006a Zhao et al. 2008).

Modified STELA

STELA was performed as described (Sfeir et al. 2005) with modifications. Ligations were performed using each of the six C-telorette oligos (T1–T6) corresponding to six permutations of the hexameric telomere repeat. Five-hundred nanogram of purified leading or lagging daughters were incubated in a 10-μL reaction (1× T4 ligase buffer, 0.001 μM individual C telorettes, 50 U New England Biolabs T4 ligase) for 20 h at 35°C. Multiple PCR amplifications were performed with 0.5 ng of ligated DNA and 0.5 μM primers (XpYpE2 forward and C-teltail reverse primers) using Abgene Hi-Fidelity PCR Master 2× mix (AB-0792, Thermo Scientific) in a final volume of 25 μL for 28 cycles of 15 sec at 94°C, 30 sec at 65°C, and 15 min at 68°C. Amplified products were resolved on a 0.7% agarose with 1× TAE and capillary-transferred onto Amersham Hybond-N + membrane overnight. Membranes were fixed for 2 h at 80°C, hybridized with a subtelomeric probe generated by PCR amplification with Xp/YpE2 and Xp/YpB2 primers, and 32 P-labeled by random priming. The membrane was then exposed to a PhosphorImager screen.

Λ Exo assay

Two micrograms of purified leading or lagging DNA was digested with 1 μL of λ exo (10 U/μL) (Epicenter Biotechnologies) for 1.5 h at 37°C. DNA was run on a 0.8% agarose gel in 1× TAE at 10 V/cm for 40 min, denatured with an alkaline solution containing 0.5 M NaOH and 1.5 M NaCl for 30 min, rinsed three times with distilled water, and dried at room temperature. The gel was then neutralized with 0.5 M Tris-HCl (pH 8) and 1.5 M NaCl for 30 min and hybridized overnight at 42°C with G-telomere probes to detect the C strands.

RNA primer pull-down assay

Prior to CsCl gradient centrifugation, genomic DNA was digested with RsaI and HaeIII, generating only blunt ends to minimize background. Two micrograms of purified leading or lagging DNA was ligated overnight at 16°C to a 0.5 μM (final concentration) mixture of the six telorette oligos (Sfeir et al. 2005) that were biotinylated. DNA was pulled down using Dynabeads kilobaseBINDER kit (Invitrogen) overnight at room temperature. DNA was released by 7.5 U of RNase HII (New England Biolabs) for 2 h. Control DNA was released using 95% formamide and 10 mM EDTA (pH 8.2) for 10 min at 90°C.


Defective telomeres are now being linked to dozens of diseases, including many types of cancer

Studying telomeres, the structures that protect the ends of chromosomes, has become a key issue in biology. In recent years, not only has their relation to ageing been confirmed defective telomeres seem to be linked to more and more illnesses, including many types of cancer. The review published by Paula Martínez and María Blasco from the CNIO in Trends in Biochemical Sciences, stresses the importance of investigating these structures to improve diagnoses and develop possible treatments for many diseases. Telomeres, in the opinion of these researchers, will become increasingly important in clinical studies.

The chromosomes in every single cell are made up of DNA and shaped like strands, with a kind of protective cap at the end of each strand of DNA. Without this end protective cap, the DNA strands would chemically bond to other strands, i.e. the chromosomes would merge and that would be lethal for the cell. The structures that prevent this catastrophe are the telomeres. They were discovered in the 1930s but decades elapsed before someone decided to study them in any depth and since the late 1990s they have always been on the cutting edge of biology research. Biologists are often surprised by their amazing and unexpected complexity, and their health-related significance.

"The biology of telomeres is extremely complex and the more we discover the more we realise what remains to be discovered," says Paula Martínez from CNIO's Telomere and Telomerase Group. "What surprises me most is the high number of factors we are finding that are essential to the preservation of telomeres and, above all, the precise coordination that is required between them all."

The fact that telomeres have been tightly preserved throughout the evolutionary tree -- in most eukaryotes: vertebrates, plants and even unicellular organisms such as yeast -- indicates their importance. In addition to preventing the merger of chromosomes, telomeres are needed to prevent the loss of genetic information each time a cell divides.

Preventing Information Loss

When a cell replicates, the molecular machinery in charge of duplicating the chromosomes -- so that each daughter cell has a copy -cannot reach the tip. This is inherently impossible due to the way the DNA replication machinery works, and it implies that any genetic material at the end of a chromosome with significant information for the cell would be lost. Telomeres prevent this from happening: they consist of a DNA sequence that does not contain genes and that is repeated numerous times- in humans and other species the sequence is TTAGGG the letters correspond to three of the building blocks that make up the DNA: thymine, adenine and guanine.

Consequently, the shortening of the DNA with every division is not significant. At least not until a certain limit is reached. When the telomeres become too short, we see the problems associated with ageing: cells reach a point where they interpret critically short telomeres as irreparable damage and react by no longer dividing, which prevents tissue from regenerating.

This happens in healthy cells but not in cancer cells. There is an enzyme, telomerase, which is capable of lengthening the telomeres de novo. This enzyme is not present in most cells of an adult organism but it is active in tumour cells. By repairing the telomeres, the telomerase enables cancer cells to proliferate and become virtually immortal.

This link to ageing and cancer, has led to the intense study of telomere-based strategies to combat cancer and diseases associated with ageing. Blasco's group has recently shown that it is possible to make cancer cells mortal by acting on the telomeres.

Zooming in to the Tip of the Buffer

The above-mentioned description of telomeres however is a simplified version of the story. We now know that there is a protective structure enveloping telomeric DNA consisting of six proteins known as shelterins, which are crucial. Another more recent discovery is that there are proteins that, although not in the telomeres themselves, interact with them at specific times to enable them to perform their functions.

These proteins enable the telomeres to unwind, for example because, the sequence repeated in telomeres, TTAGGG, ends in a single strand of DNA that curves forming a loop and connects to the original strand of the double chain forming a triple chain. "Yes, it is very complicated," admits Martínez. "Structures of up to four chains of DNA can form."

When a cell divides, the telomeres are also replicated. This implies that the end loop must unwind first and then form again. This process also contributes to the shortening of telomeres and we now know that some of the shelterins as well as other associated proteins that interact with telomeres are key elements in this process.

Telomere Syndromes

According to Martínez, "there is now more evidence about relationship between telomere maintenance and several illnesses."

Telomere syndromes, or telomeropathies, have been identified in patients with mutations of the telomerase enzyme. This group includes, for example, pulmonary fibrosis and problems related to the malfunction of the bone marrow. A direct relationship between telomere dysfunctions and many types of cancer has also been found. More recently, we have also discovered that mutations of the proteins that protect telomeric DNA, the shelterins, and those that interact with the telomeres, are linked to various diseases, such as dyskeratosis congenita, Hoyeraal-Hreidarsson syndrome or Revesz syndrome.

"These discoveries underline the plethora of components and pathways that control telomere functions," write the authors in the paper. "In the future, research will bring to light more unknown factors that will improve our understanding of the mechanisms governing cancer and syndromes linked to the shortening of telomeres. We hope that this knowledge will be transferred to the clinic in order to improve the diagnosis and treatment of diseases."


Why do telomeres shorten during replication?

I'm curious why this happens. It seems that the rest of the dna is copied in full, barring teeny mistakes, so why aren't Telomeres?

This is a problem called the "end-replication problem". This relies on fundamentally understanding that while DNA is bidirectional (each complementary strand has a 5' to 3' sequence), the DNA polymerase that replicates the daughter strands is unidirectional (synthesizing only from the 5' to 3' end).

As a result, daughter strand synthesis occurs differently between the 3'-5' mother and the 5'-3' mother. The 3'-5' mother is called the leading strand, and an RNA primer is able to anneal to the 3' end of the mother, and allow for DNA polymerase to replicate from the primer to the very end (since the daughter will be running from the 5'-3' in the direction of the DNA polymerase).

However, the 5'-3' mother is problematic. This strand is the lagging strand, in which multiple RNA primers must be used to create discontinuous 'Okazaki fragments" of DNA as helicase unzips the mother double helix. Normally the RNA primers are replaced and filled in with DNA polymerase and ligated upon complete unzipping. However, the final RNA primer on the 3' end of the mother strand cannot be filled in with DNA, and the RNA primer is degraded.

As a result, as DNA is replicated over and over, approximately 50-200bp of DNA is unreplicated at the mother 3' end. Since the ends are capped with telomeres, telomere shortening inevitably occurs.

In theory, telomerase can act to ɿill-in' this gap by capping the unreplicated end with fresh telomeres. However, telomerase activity is NOT expressed by most somatic cells, and primarily functions in germ and some embryonic cells.


Recombinogenic Telomeres in Diploid Sorex granarius (Soricidae, Eulipotyphla) Fibroblast Cells

FIG 1 Position of centromeres and nucleolar relationship of acrocentric chromosomes in primary S. granarius fibroblasts. (A) Centromere position on S. granarius acrocentric chromosomes. A two-step experiment was performed, including immunofluorescence assay with ANA-CREST antibodies to detect centromeric proteins (green) and subsequent two-color FISH with a C-rich telomeric PNA probe (Cy3 red) and an 18S rDNA probe (pseudocolored blue). Chromosomes were counterstained with DAPI (pseudocolored white). Centromeres are generally found adjacent to or overlapping signals from telomeric and rDNA probes. (B) Visualization of transcriptionally active nucleoli in primary S. granarius fibroblasts by use of anti-UBF1 antibodies (red). Telomeres were subsequently visualized through PNA FISH (green). In the optical slice at left, arrows indicate close contacts between telomeres and nucleoli. The mean number of nuclear telomere foci (± SEM) was 13.7 ± 2.8 (range, 9 to 26), with 65% ± 2.6% of them being in association with UBF1 signals (n = 90 nuclei). Bar, 10 μm.

The telomere-associated rDNA loci in S. granarius are not always expressed.

S. granarius fibroblasts maintain telomere function after prolonged in vitro culture.

FIG 2 Chromosome stability in long-term-culture S. granarius fibroblast cells. (A) Representative karyotype of S. granarius fibroblasts in long-term culture (passage 132 approximately 1 1/2 years). The diploid chromosome complement in female S. granarius is 36, with 32 acrocentric chromosomes and 4 metacentric chromosomes. Arrow, additional chromosome “r.” (B) Chromosome number variation in late-passage fibroblasts (passage 132). Two hundred metaphase spreads were analyzed. (C) Representative images of telomere length analysis by Q-FISH on metaphase chromosomes from primary (passage 7 p7) and long-term (passage 116 p116) fibroblast cultures. The telomeric PNA C-rich probe (Cy3 red) was used, and chromosomes were counterstained with DAPI. The telomere length inequality reported previously for primary fibroblasts (long telomeres on short arms of acrocentric chromosomes versus very short telomeres on all other extremities [10]) was preserved in long-term cultures. Bars, 10 μm. (D) Quantification of Q-FISH telomere intensities for long and short telomeres. See Table 1 for more details.
Telomeres and passageNo. of telomeres analyzedMean fluorescence intensity ± SEM
Long
733388,392.5 ± 1,200.7
11634281,593.8 ± 2,613.9 *
Short
77852,718.7 ± 69.1
1167941,494.3 ± 111.2 *

S. granarius fibroblasts are telomerase positive.

FIG 3 Fibroblasts of S. granarius contain active telomerase. (A and B) Conventional TRAP analysis of S. granarius fibroblast cells revealed telomerase activity in primary and long-term-culture cells. (A) DNA PAGE gel after conventional TRAP using protein extracts from S. granarius fibroblasts at different passages. A protein extract from PHA-stimulated human leukocytes was used as a positive control. (B) Densitometric analysis of conventional TRAP results. (C) A real-time semiquantitative TRAP assay was performed on cell extracts prepared from early-passage S. granarius cells (passage 9) and compared to the activity found in an equal number of human HeLa cells. Serial dilutions of S. granarius whole-cell extracts (3,000, 1,000, and 300 cell equivalents) resulted in a decreased TRAP activity. Treatment with RNase A and omission of the cell extract served as negative controls.

S. granarius fibroblasts display ALT-associated PML-like bodies.

FIG 4 S. granarius fibroblasts carry APB-like structures. (A) S. granarius primary fibroblasts (passage 7) were transfected with a plasmid expressing an ECFP-ICP0* fusion to reveal putative PML nuclear bodies. The native fluorescence of the ECFP-ICP0* fusion in three transfected nuclei is shown. ICP0* formed 7 to 24 (18 ± 1.94 [mean ± SEM]) large nuclear foci per transfected nucleus (n = 150 nuclei). (B) Cells expressing ECFP-ICP0* were costained with antibodies against the human shelterin protein RAP1 (green) and with a telomeric C-rich PNA probe (Cy3 red) and mounted in Vectashield with DAPI. Maximum-intensity projections are presented for all color channels. The arrows point to close contacts between RAP1, telomeres, and ICP0* (readily visible in the merged image on the far right), suggesting that they are part of the same APB-like structure. (C) 3D reconstitution of the images in panel B. (D) Immuno-FISH on S. granarius metaphase chromosomes, combining antibodies to human RAP1 (green) and a telomeric PNA probe (Cy3 red) chromosomes are stained with DAPI. Bars, 10 μm.

S. granarius telomeres show active recombination.

FIG 5 Recombination at telomeres in S. granarius . (A) CO-FISH approach to reveal T-SCEs. After the removal of newly synthesized strands, a C-rich probe will exclusively detect the parental G-rich strand (one-color CO-FISH). If an exchange has taken place after replication, two signals instead of one will be detected at the chromosome extremity. In two-color CO-FISH, strand-specific C-rich and G-rich telomeric probes are used, and T-SCEs are detected as mixed red-green signals. (B) CO-FISH using two strand-specific telomeric probes: TelPNA-C-rich-Cy3 (red) and Tel-LNA-G-rich-FAM (green). Chromosomes were counterstained with DAPI (blue). Only long telomeres present on short acrocentric arms were analyzed. Most extremities show one single green or red robust signal per chromatid. The box indicates a chromosome with mixed signals, indicating a T-SCE. Affected chromosomes varied from metaphase to metaphase. Quantifications of different experiments using one- or two-color CO-FISH are presented in panel E and Table 2. (C) Examples of T-SCEs detected in S. granarius early-passage fibroblasts (p15). (D) Signal enhancement allows the detection of potential highly asymmetric exchanges (very weak green signals colocalizing with the strong red signal, and vice versa [arrowheads]). Enlarged examples are presented. The segregated CO-FISH analysis presented in Fig. 6 indicates that such colocalizations correspond to bona fide T-SCEs. Bars, 10 μm. (E) Quantification of T-SCEs in S. granarius early-passage (S. gr p15) fibroblasts relative to U2OS/ALT and HT1080/TEL + human cancer cell lines (n = 30 metaphase spreads for each condition). The frequency of metaphase chromosomes carrying T-SCEs when only “robust” fluorescence signals are taken into account appears to be low in S. granarius fibroblasts. However, when T-SCEs are searched upon enhancement of signals, the frequency is much higher. The fact that these are bone fide T-SCEs was confirmed by segregated CO-FISH analysis (Fig. 6).
Culture type and expt no.No. of metaphase spreads analyzedNo. of extremities with long telomeres No. of metaphase spreads with: % of metaphase spreads with T-SCEs (±SEM)
1 T-SCE>1 T-SCE
Asynchronous
1862,73840552.3
2902,87043755.6
3702,21637458.6
Total2467,8241201655.5 (±3.8)
Synchronous
11003,19142749.0
2902,87243856.6
Total1906,063851552.3 (±4.8)
FIG 6 (A) BrdU/C incorporation during two cell cycles prior to the CO-FISH procedure, using strand-specific telomeric probes, results in segregation of the unsubstituted G- and C-rich strands into different chromosomes, such that during the second M phase, every chromosome extremity will be stained, after the CO-FISH procedure, with only one probe, either red (TelPNA-C-rich-Cy3) or green (TelLNA-G-rich-FAM). (B) If a T-SCE occurs during the first cell cycle, the two unsubstituted strands will cosegregate during the first mitosis and will be detected on different sister chromatids of the same chromosome during the second M phase. The CO-FISH procedure will then reveal one red and one green signal on the same chromosome extremity (marked 1). If a T-SCE occurs during the second cell cycle, this exchange will result in same-color doublets, either red or green (marked 2). However, if a second exchange affects an extremity that had already undergone T-SCE during the first cell cycle, doublets will be of both colors (marked 1&2). (C) Two-color segregated CO-FISH in S. granarius . Highly asymmetric two-color doublets are frequently detected in S. granarius fibroblasts (p15). Examples of such T-SCEs are enlarged and color decomposed on the right. The TelPNA-C-rich-Cy3 probe is more efficient than the TelLNA-G-rich-FAM probe for detecting small doublets. Chromosomes were counterstained with DAPI (blue). Bar, 10 μm. FIG 7 Telomeric circles are detected in primary S. granarius fibroblasts. Ten micrograms of genomic DNA from S. granarius early-passage (p13) fibroblasts (A) and 20 μg genomic DNA from the human ALT cancer cell line U2OS (B) were digested with MboI, separated by 2D gel electrophoresis, transferred onto N+ nylon membranes, and hybridized with a digoxigenin-labeled telomeric C-rich oligonucleotide. Arrow 1, single-stranded linear DNA arrow 2, double-stranded linear DNA arrow 3, circular DNA.

S. granarius telomeres bear marks of DNA damage.

FIG 8 Spontaneous telomere dysfunction in primary S. granarius fibroblasts. (A) Meta-TIF analysis of S. granarius fibroblasts (one-step protocol). Metaphase chromosomes were first stained with anti-γH2AX antibodies (green) and subsequently stained with a telomeric PNA probe (red) for detection of telomere-induced foci (TIF). Chromosomes were counterstained with DAPI (blue). (B) Relative green and red fluorescence intensities of particular chromosomes from the metaphase spread shown in panel A, illustrating either perfect colocalization of red and green signals or the spread of green signals toward the interstitial region. (C) Meta-TIF analysis of S. granarius fibroblasts by a two-step protocol involving immunofluorescence assay with anti-γH2AX antibodies, with image acquisition (green left panel), as well as hybridization with a telomeric PNA probe, with visualization (red middle panel). The right panel show the merge of the two visualization steps. (D) Illustration of chromosome- and chromatid-type TIF detected in a two-step experiment. The images show staining with anti-γH2AX antibodies (green) and a telomeric PNA probe (red). A quantification of these experiments is presented in Table 3. (E) Detection of TIF in S. granarius interphase nucleus by a one-step protocol. The image shows staining with anti-γH2AX antibodies (green) and a telomeric PNA probe (red). Bars, 10 μm.
Expt (n)No. of metaphase spreads analyzedNo. of chromosomes analyzedAvg no. of chromosomes per metaphase spread Avg no. of DDR + telomeres per metaphase spread (±SEM)
Long telomeresShort telomeres
One-step expt (2)1003,58835.914.95 (±3.05)NA
Two-step expt (1)622,22835.922 (±2.32), among which 16.3 (±4.5) were of the chromosome type10 (±1.67), among which 7.3 (±1.8) were of the chromosome type

Very short telomeres accumulate upon antitelomerase treatment of S. granarius fibroblasts.

S. granarius telomeres are transcribed.

FIG 9 Both G-rich and C-rich telomere strands are expressed in S. granarius cells. (A) RNA FISH experiments with strand-specific probes on S. granarius interphase nuclei. The C-rich probe detected either the UUAGGG or TTAGGG sequence, and the G-rich probe detected either CCCUAA or CCCTAA, depending on whether the hybridization was done on native or denatured nuclei, respectively. Signals were visible with both probes under native conditions without RNase treatment, albeit at lower intensity when the probe was G-rich. The signal was completely lost when native preparations were treated with RNase. Both probes detected telomeres equally efficiently under denaturing conditions. Nuclei are stained with DAPI (blue). (B) Similar RNA FISH experiment on metaphase chromosome preparations obtained by cytospin centrifugation under native or denatured conditions and treated or not treated with RNase. Once again, the C-rich probe yielded stronger signals than the G-rich probe, only on native, RNase-untreated chromosomes. Bars, 10 μm. (C) Northern blot analysis of S. granarius telomeric RNA transcripts in cells at different passages (p). (Left) Visualization of total RNA in the gel by ethidium bromide (EtBr) staining before transfer onto a membrane. (Second panel) Hybridization with a C-rich telomere probe labeled with digoxigenin. (Third panel) Verification of the stripping efficiency. (Right) Hybridization with a G-rich probe labeled with digoxigenin.

DNA Replication

When a cell divides, it is important that each daughter cell receives an identical copy of the DNA. This is accomplished by the process of DNA replication. The replication of DNA occurs during the synthesis phase, or S phase, of the cell cycle, before the cell enters mitosis or meiosis.

The elucidation of the structure of the double helix provided a hint as to how DNA is copied. Recall that adenine nucleotides pair with thymine nucleotides, and cytosine with guanine. This means that the two strands are complementary to each other. For example, a strand of DNA with a nucleotide sequence of AGTCATGA will have a complementary strand with the sequence TCAGTACT ([link]).

Because of the complementarity of the two strands, having one strand means that it is possible to recreate the other strand. This model for replication suggests that the two strands of the double helix separate during replication, and each strand serves as a template from which the new complementary strand is copied ([link]).

During DNA replication, each of the two strands that make up the double helix serves as a template from which new strands are copied. The new strand will be complementary to the parental or “old” strand. Each new double strand consists of one parental strand and one new daughter strand. This is known as semiconservative replication. When two DNA copies are formed, they have an identical sequence of nucleotide bases and are divided equally into two daughter cells.

DNA Replication in Eukaryotes

Because eukaryotic genomes are very complex, DNA replication is a very complicated process that involves several enzymes and other proteins. It occurs in three main stages: initiation, elongation, and termination.

Recall that eukaryotic DNA is bound to proteins known as histones to form structures called nucleosomes. During initiation, the DNA is made accessible to the proteins and enzymes involved in the replication process. How does the replication machinery know where on the DNA double helix to begin? It turns out that there are specific nucleotide sequences called origins of replication at which replication begins. Certain proteins bind to the origin of replication while an enzyme called helicase unwinds and opens up the DNA helix. As the DNA opens up, Y-shaped structures called replication forks are formed ([link]). Two replication forks are formed at the origin of replication, and these get extended in both directions as replication proceeds. There are multiple origins of replication on the eukaryotic chromosome, such that replication can occur simultaneously from several places in the genome.

During elongation, an enzyme called DNA polymerase adds DNA nucleotides to the 3' end of the template. Because DNA polymerase can only add new nucleotides at the end of a backbone, a primer sequence, which provides this starting point, is added with complementary RNA nucleotides. This primer is removed later, and the nucleotides are replaced with DNA nucleotides. One strand, which is complementary to the parental DNA strand, is synthesized continuously toward the replication fork so the polymerase can add nucleotides in this direction. This continuously synthesized strand is known as the leading strand. Because DNA polymerase can only synthesize DNA in a 5' to 3' direction, the other new strand is put together in short pieces called Okazaki fragments. The Okazaki fragments each require a primer made of RNA to start the synthesis. The strand with the Okazaki fragments is known as the lagging strand. As synthesis proceeds, an enzyme removes the RNA primer, which is then replaced with DNA nucleotides, and the gaps between fragments are sealed by an enzyme called DNA ligase.

The process of DNA replication can be summarized as follows:

  1. DNA unwinds at the origin of replication.
  2. New bases are added to the complementary parental strands. One new strand is made continuously, while the other strand is made in pieces.
  3. Primers are removed, new DNA nucleotides are put in place of the primers and the backbone is sealed by DNA ligase.

You isolate a cell strain in which the joining together of Okazaki fragments is impaired and suspect that a mutation has occurred in an enzyme found at the replication fork. Which enzyme is most likely to be mutated?

Telomere Replication

Because eukaryotic chromosomes are linear, DNA replication comes to the end of a line in eukaryotic chromosomes. As you have learned, the DNA polymerase enzyme can add nucleotides in only one direction. In the leading strand, synthesis continues until the end of the chromosome is reached however, on the lagging strand there is no place for a primer to be made for the DNA fragment to be copied at the end of the chromosome. This presents a problem for the cell because the ends remain unpaired, and over time these ends get progressively shorter as cells continue to divide. The ends of the linear chromosomes are known as telomeres, which have repetitive sequences that do not code for a particular gene. As a consequence, it is telomeres that are shortened with each round of DNA replication instead of genes. For example, in humans, a six base-pair sequence, TTAGGG, is repeated 100 to 1000 times. The discovery of the enzyme telomerase ([link]) helped in the understanding of how chromosome ends are maintained. The telomerase attaches to the end of the chromosome, and complementary bases to the RNA template are added on the end of the DNA strand. Once the lagging strand template is sufficiently elongated, DNA polymerase can now add nucleotides that are complementary to the ends of the chromosomes. Thus, the ends of the chromosomes are replicated.

Telomerase is typically found to be active in germ cells, adult stem cells, and some cancer cells. For her discovery of telomerase and its action, Elizabeth Blackburn ([link]) received the Nobel Prize for Medicine and Physiology in 2009.

Telomerase is not active in adult somatic cells. Adult somatic cells that undergo cell division continue to have their telomeres shortened. This essentially means that telomere shortening is associated with aging. In 2010, scientists found that telomerase can reverse some age-related conditions in mice, and this may have potential in regenerative medicine. 1 Telomerase-deficient mice were used in these studies these mice have tissue atrophy, stem-cell depletion, organ system failure, and impaired tissue injury responses. Telomerase reactivation in these mice caused extension of telomeres, reduced DNA damage, reversed neurodegeneration, and improved functioning of the testes, spleen, and intestines. Thus, telomere reactivation may have potential for treating age-related diseases in humans.

DNA Replication in Prokaryotes

Recall that the prokaryotic chromosome is a circular molecule with a less extensive coiling structure than eukaryotic chromosomes. The eukaryotic chromosome is linear and highly coiled around proteins. While there are many similarities in the DNA replication process, these structural differences necessitate some differences in the DNA replication process in these two life forms.

DNA replication has been extremely well-studied in prokaryotes, primarily because of the small size of the genome and large number of variants available. Escherichia coli has 4.6 million base pairs in a single circular chromosome, and all of it gets replicated in approximately 42 minutes, starting from a single origin of replication and proceeding around the chromosome in both directions. This means that approximately 1000 nucleotides are added per second. The process is much more rapid than in eukaryotes. [link] summarizes the differences between prokaryotic and eukaryotic replications.

Differences between Prokaryotic and Eukaryotic Replications
Property Prokaryotes Eukaryotes
Origin of replication Single Multiple
Rate of replication 1000 nucleotides/s 50 to 100 nucleotides/s
Chromosome structure circular linear
Telomerase Not present Present

Click through a tutorial on DNA replication.

DNA Repair

DNA polymerase can make mistakes while adding nucleotides. It edits the DNA by proofreading every newly added base. Incorrect bases are removed and replaced by the correct base, and then polymerization continues ([link]a). Most mistakes are corrected during replication, although when this does not happen, the mismatch repair mechanism is employed. Mismatch repair enzymes recognize the wrongly incorporated base and excise it from the DNA, replacing it with the correct base ([link]b). In yet another type of repair, nucleotide excision repair, the DNA double strand is unwound and separated, the incorrect bases are removed along with a few bases on the 5' and 3' end, and these are replaced by copying the template with the help of DNA polymerase ([link]c). Nucleotide excision repair is particularly important in correcting thymine dimers, which are primarily caused by ultraviolet light. In a thymine dimer, two thymine nucleotides adjacent to each other on one strand are covalently bonded to each other rather than their complementary bases. If the dimer is not removed and repaired it will lead to a mutation. Individuals with flaws in their nucleotide excision repair genes show extreme sensitivity to sunlight and develop skin cancers early in life.

Most mistakes are corrected if they are not, they may result in a mutation—defined as a permanent change in the DNA sequence. Mutations in repair genes may lead to serious consequences like cancer.

Section Summary

DNA replicates by a semi-conservative method in which each of the two parental DNA strands act as a template for new DNA to be synthesized. After replication, each DNA has one parental or “old” strand, and one daughter or “new” strand.

Replication in eukaryotes starts at multiple origins of replication, while replication in prokaryotes starts from a single origin of replication. The DNA is opened with enzymes, resulting in the formation of the replication fork. Primase synthesizes an RNA primer to initiate synthesis by DNA polymerase, which can add nucleotides in only one direction. One strand is synthesized continuously in the direction of the replication fork this is called the leading strand. The other strand is synthesized in a direction away from the replication fork, in short stretches of DNA known as Okazaki fragments. This strand is known as the lagging strand. Once replication is completed, the RNA primers are replaced by DNA nucleotides and the DNA is sealed with DNA ligase.

The ends of eukaryotic chromosomes pose a problem, as polymerase is unable to extend them without a primer. Telomerase, an enzyme with an inbuilt RNA template, extends the ends by copying the RNA template and extending one end of the chromosome. DNA polymerase can then extend the DNA using the primer. In this way, the ends of the chromosomes are protected. Cells have mechanisms for repairing DNA when it becomes damaged or errors are made in replication. These mechanisms include mismatch repair to replace nucleotides that are paired with a non-complementary base and nucleotide excision repair, which removes bases that are damaged such as thymine dimers.

Art Connections

[link] You isolate a cell strain in which the joining together of Okazaki fragments is impaired and suspect that a mutation has occurred in an enzyme found at the replication fork. Which enzyme is most likely to be mutated?


Calorie Restriction

Some studies have shown that calorie restriction (i.e., a 20-40% reduction of dietary caloric intake) extends life expectancy in several species ranging from yeast to rodents. One possible explanation is that caloric restriction reduced production of reactive oxygen species by mitochondria. In addition, calorie restriction induces autophagy that removes harmful proteins and organelles, thereby reducing the accumulating damage to the cell. However, this phenomenon has not yet been demonstrated in humans. For an excellent review, see Ivanova DG, Yankova TM: The free radical theory of aging in search of a strategy for increasing life span. Folia Med 201355(1):33-41.

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Date last modified: December 14, 2013.
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