Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression

Catarina Salgado; Celine Roelse; Rogier Nell; Nelleke Gruis; Remco van Doorn; Pieter van der Velden

doi:10.1371/journal.pone.0231418

. 2020 Apr 8;15(4):e0231418. doi: 10.1371/journal.pone.0231418

Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression

Catarina Salgado ¹, Celine Roelse ¹, Rogier Nell ², Nelleke Gruis ¹, Remco van Doorn ¹, Pieter van der Velden ^2,^*

Editor: Srinivas Saladi³

PMCID: PMC7141627 PMID: 32267900

Abstract

The telomerase reverse transcriptase (TERT) gene is responsible for telomere maintenance in germline and stem cells, and is re-expressed in 90% of human cancers. CpG methylation in the TERT promoter (TERTp) was correlated with TERT mRNA expression. Furthermore, two hotspot mutations in TERTp, dubbed C228T and C250T, have been revealed to facilitate binding of transcription factor ETS/TCF and subsequent TERT expression. This study aimed to elucidate the combined contribution of epigenetic (promoter methylation and chromatin accessibility) and genetic (promoter mutations) mechanisms in regulating TERT gene expression in healthy skin samples and in melanoma cell lines (n = 61). We unexpectedly observed that the methylation of TERTp was as high in a subset of healthy skin cells, mainly keratinocytes, as in cutaneous melanoma cell lines. In spite of the high promoter methylation fraction in wild-type (WT) samples, TERT mRNA was only expressed in the melanoma cell lines with either high methylation or intermediate methylation in combination with TERT mutations. TERTp methylation was positively correlated with chromatin accessibility and TERT mRNA expression in 8 melanoma cell lines. Cooperation between epigenetic and genetic mechanisms were best observed in heterozygous mutant cell lines as chromosome accessibility preferentially concerned the mutant allele. Combined, these results suggest a complex model in which TERT expression requires either a widely open chromatin state in TERTp-WT samples due to high methylation throughout the promoter or a combination of moderate methylation fraction/chromatin accessibility in the presence of the C228T or C250T mutations.

Introduction

Approximately 90% of all human cancers share a transcriptional alteration: reactivation of the telomerase reverse transcriptase (TERT) gene [1, 2]. TERT encodes the catalytic subunit of the ribonucleoprotein telomerase and is capable of extending the repetitive, non-coding DNA sequence on terminal ends of chromosomes, the telomeres. As the single-stranded 5’ ends of chromosomes are shortened with each cellular division, telomeres prevent loss of coding chromosomal DNA [3–6]. Telomerase is only transcribed in a subset of stem cells in growing or renewing tissues, but through reactivation of telomerase expression, cells can extend telomeres or prevent telomeres shrinkage. This is termed telomere maintenance, which is one of the hallmarks of cancer, and allows subsequent indefinite proliferation and immortalization [3, 6–8].

Since the MYC oncogene has firstly been identified to activate telomerase, a variety of epigenetic or genetic mechanisms in the gene body or TERT promoter (TERTp) have followed, such as CpG methylation, histone modifications, mutations, germline genetic variations, structural variations, DNA amplification or chromosomal rearrangements [3, 5, 7].

A widely investigated mechanism that could induce TERT reactivation is the presence of mutations in the gene promoter [7, 9]. Horn and Huang et al. identified two mutually exclusive TERTp point mutations that are correlated to TERT mRNA expression by creating binding motifs for the transcription factor E26 transformation-specific/ternary complex factor (ETS/TCF) [7, 9]. These mutations, chr5:1,295,228 C>T and chr5:1,295,250 C>T in hg19 (−124 bp and −146 bp from the translation start site, respectively), henceforth respectively dubbed C228T and C250T, were first identified in melanoma. Furthermore, these mutations showed high prevalence in and were correlated with poor prognosis of cutaneous melanomas [4, 5, 10–12].

An additional mechanism by which a gene can be made accessible to transcription factors, facilitating gene expression, is hypomethylation of promoter CpG islands, a hallmark of euchromatin [13, 14]. Methylation located in the gene body, however, shows a positive correlation with active gene expression [15]. In stark contrast to most genes, TERTp hypermethylation may also allow gene expression since transcriptional repressors rely on unmethylated promoter CpGs, such as CCCTC-binding factor (CTCF)/cohesin complex or MAZ [16–18]. As such, in combination with transcription factor binding, dissociation of the repressor may result in TERT expression [3, 16, 19, 20]. Castelo-Branco et al. proposed that methylation of a specific CpG site in TERTp, cg11625005 (position 1,295,737 in hg19) was associated with paediatric brain tumours progression and poor prognosis [20]. This finding was later supported by the study from Barthel et al., in which the CpG methylation was found to be correlated with TERT expression in samples lacking somatic TERT alterations and to be generally absent in normal samples adjacent to tumour tissue [3].

Chromatin organisation, its plasticity and dynamics at TERTp region have been reported as relevant players in regulation of gene expression by influencing the binding of transcription factors [21, 22]. Cancer cells are positively selected to escape the native repressive chromatin environment in order to allow TERT transcription [23].

In the present study, we aim to elucidate the interaction of genetic and epigenetic mechanisms in regulation of TERTp. We approach this by using novel droplet digital PCR (ddPCR)-based assays [24]. Human-derived benign skin cells (keratinocytes, dermal fibroblasts, melanocytes, skin biopsy samples and naevi) and melanoma cell lines were analysed. The TERTp mutational status was assessed along with the absolute presence of methylation in the TERTp at a CpG-specific resolution. The effect of chromatin accessibility in TERT expression was evaluated in a subset of cultured melanoma cell lines.

Results

NGS-based deep bisulfite sequencing and development of a ddPCR assay to assess TERTp methylation fraction

We first aimed to quantitatively measure the TERTp methylation at a CpG-specific resolution in primary skin samples and melanoma cell lines. DNA of 44 primary skin biopsy samples and melanoma cell lines was bisulfite-converted (BC) and analysed using NGS-based deep bisulfite sequencing to assess the methylation fraction (MF) in a region of TERTp encompassing 31 CpG sites. The TERTp MF was high in some healthy skin samples, such as normal skin (~30%), naevi (~30%) and cultured keratinocytes (~50%). In the latter group, in fact, the MF was as high as in cutaneous melanoma cell lines (Figs 1 and 7A). In contrast, the fibroblasts and low-passage cultured melanocytes show the lowest MF observed in this cohort. Since the cutaneous melanoma originates from melanocytes of the skin, we found the difference in MF between normal melanocytes and cutaneous melanoma cells quite remarkable.

Fig 1 — DNA samples were bisulfite-converted (BC) and analysed through NGS-based deep sequencing. Connected scatter plot representing the MF per cell type group in absolute distance between measured CpG sites. Blue arrow: cg11625005 (position 1,295,737). Samples included: fibroblasts (n = 5), melanocytes (n = 5), naevi (n = 6), normal skin samples (n = 11), keratinocytes (n = 8), cutaneous melanoma cell lines (n = 6) and uveal melanoma cell lines (n = 3).

Fig 7 — Schematic representation of *TERT*p with the relative positions of cg11625005 (position 1,295,737 in hg19) to the *TERT*p mutations (position 1,295,228 and 1,295,250) and the transcription start site (TSS). A. Heat-map of methylation fraction (MF) in 31 CpG sites (top) in 44 samples (left). Yellow-marked CpG cg11625005 (position 1,295,737) is recognised by MSRE HgaI. Blue-marked CpG in 1,295,731 is recognised by MSRE AvaI. Black rectangle: MF at the cg11625005 measured either by NGS (clear squares, n = 44) and by ddPCR (patterned squares, n = 17; these samples were not included in the 44-sample batch subjected to NGS). B. *TERT* mRNA expression in 31 samples by qPCR analysed through the ΔΔCT method in Bio-Rad CFX manager software (version 3.1, Bio-Rad). C. *TERT*p mutations evaluated through ddPCR with commercial *TERT* C250T and C228T Mutation Assays in total 61 samples. D. Analysis of the chromatin accessibility in 8 cultured cell lines for *TERT* methylation region using *GAPDH* as a positive control.

In order to validate the TERTp MF obtained through NGS in a quantitative manner, we have developed a ddPCR assay (Fig 2A) using methylation-sensitive restriction enzymes (MSREs) HgaI and AvaI, which recognise the CpG on position 1,295,737 (cg11625005) and 1,295,731 in hg19, respectively. Castelo-Branco et al. showed that methylation of the cg11625005 in TERTp, was associated with tumour progression and poor prognosis of childhood brain tumours [20]. Barthel et al. affirmed a correlation between methylation and TERT expression in samples lacking somatic TERT alterations and a lower methylation level in normal samples [3]. Indeed, in our study, the MF of fibroblasts was as low as that of the unmethylated control DNA, whereas that of the keratinocytes was higher than most of the cutaneous melanoma cell lines (Fig 2B). The MF of cg11625005 (position 1,295,737) obtained through NGS and by ddPCR were highly correlated (R² = 0.82, p<0.001) (Fig 2C). The MF of 1,295,731 assessed through ddPCR even yielded a stronger correlation (R² = 0.96, p<0.001) (Fig 2D).

Absence of correlation between methylation fraction and TERT expression

Cancer cells are commonly characterised by hypermethylation of promoter CpG islands resulting in repression of tumour suppressor genes. However, in TERT, promoter hypermethylation was found to be associated with higher expression, since CTCF repressors of TERT transcription do not bind methylated sequences [3, 16, 17, 19]. In our sample cohort, there was no correlation between TERT methylation of cg11625005 and mRNA expression (n = 31, Figs 3 and 7B).

Fig 3 — Linear regression and correlation analysis were performed (F_(1,29) = 1.13, r = 0.19, ns p = 0.297).

Evaluation of TERTp mutations in a collection of skin samples and melanoma cell lines

Besides promoter methylation, somatic mutations are also known to be correlated with TERTp reactivation. Therefore, we characterised the TERTp mutational status of the sample cohort. Sanger sequencing on one naevus, fresh skin and cutaneous melanoma cell lines 518A2, 607B, A375, 94.07 and 93.08 revealed melanoma-associated TERT C250T and C228T mutations (Fig 4A). Aiming to use the ddPCR method to evaluate the mutational load of the samples, the TERT C250T and C228T mutation assays were validated in three samples of which the mutation was identified in sequencing analysis, 518A2, 607B and A375 (Fig 4B). Following the test runs, the C228T and C250T assays were used on the extended sample cohort (n = 61) (S5 Table and Fig 7C). All TERTp-mutated samples were cutaneous melanoma cell lines, however OCM8 and 94.13 cutaneous cell lines tested wild-type. The C250T mutation was not present in combination with the C228T mutation in any sample, confirming that the mutations are mutually exclusive.

Fig 4 — A. The *TERT*p region encompassing the C228T and C250T mutations was sequenced through Sanger sequencing using McEvoy’s [25] *TERT*p forward primer. The *TERT*p region of fresh skin 1, Naevus 1, 518A2, 607B, A375, 94.07, 93.08 is shown. The left and right arrows respectively indicate the positions 1,295,228 and 1,295,250. R: one-letter code for bases G or A; Green arrow: wild-type; red arrow: C>T mutation on the complementary strand. B. Evaluation of *TERT*p mutations through commercial Bio-Rad TERT assays in 518A2, 607B and A375 melanoma cell lines. 2D ddPCR plots of the results from the C228T mutation assay (left) and C250T mutation assay (right). The blue cloud represents mutant copies; the green cloud represents WT copies.

Absence of correlation between mutational status and TERT expression

As the presence of mutations in the gene promoter induces TERT reactivation, we assessed the correlation between mutational status with TERT mRNA expression (n = 31). When WT and mutated samples (either C228T or C250T) were compared, regardless of origin of the tissue, no significant differences for TERT mRNA expression were found (Fig 5). Moreover, TERT expression was exclusive to the melanoma cell lines, either with or without TERTp mutations (Fig 7B).

Fig 5 — One-way ANOVA (F_(2,28) = 1.75, ns p = 0.192).

TERT expression is correlated to chromatin accessibility

In contrast to most genes, methylation of the TERTp positively correlates with its mRNA expression [3, 16, 17, 19]. Although we were not able to confirm this finding, we investigated whether besides promoter methylation, other mechanisms could contribute to chromatin accessibility to transcription factors affecting TERTp regulation. Therefore, we analysed chromatin state in a subset of melanoma cell lines (cutaneous, 518A2, 607B, 94.07, A375, 93.08 and OCM8; and uveal, OMM2.5 and Mel270) by ddPCR methodology instead of qPCR for an accurate quantification. The positive control gene GAPDH, a housekeeping gene that is generally expressed in all conditions, and thus 100% accessible, was used. The accessibility in the region around cg11625005 shows a high variability, being over 90% in uveal cell lines while being intermediate to low in cutaneous melanoma cell lines (Figs 6A and 7D and S6 Table). When comparing the accessibility around cg11625005 to the methylation fraction of this CpG, a significant positive correlation was observed (R² = 0.89, p<0.001) (Fig 6B). Another positive correlation (R² = 0.59, p<0.05) was found when comparing the accessibility of the same region to the normalised TERT mRNA expression levels in these samples (Fig 6C). In actuality, in this subset of 8 cell lines, the TERTp methylation and gene expression show a statistically significant (p<0.05) positive correlation (Fig 6D). The 3 cell lines with higher MF are those with the highest chromatin accessibility (OMM2.5, Mel270 and OCM8). Remarkably, these are also the cell lines with WT-TERTp, in which the chromatin accessibility was significantly higher than in the mutated subgroup (Fig 6E).

Fig 6 — Cell lines (518A2, 607B, 94.07, A375, 93.08, OMM2.5, Mel270 and OCM8) were analysed with the EpiQ chromatin kit, and ddPCR was performed using primers and probes for positive control gene *GAPDH* and for the *TERT* methylation region, a 231-bp amplicon around cg11625005. Accessibility (%) was calculated by the ratio of the digested sample to its matched undigested sample, subtracted from 1, and subsequently normalised against the positive control *GAPDH*. A. Accessibility of the *TERT* methylation region relative to *GAPDH* (mean ± SD, multiple t-tests, one t-test per cell line, *p<0.001, ns p = 0.149). B & C. Correlation plots of gene accessibility around cg11625005 with the MF (%) of cg11625005 obtained through ddPCR (B), or with normalised expression levels via qPCR (C). Linear regression and correlation analysis were performed (F_(1.6) = 49.9, r = 0.95, p<0.001 and F_(1,6) = 8.6, r = 0.77, p<0.05, respectively). D. Correlation plot between MF (%) of cg11625005 obtained through ddPCR and normalised expression levels via qPCR. Linear regression and correlation analysis were performed (F_(1.6) = 16.92, r = 0.86, p<0.05). E. Comparison of WT (OMM2.5, Mel270 and OCM8) and mutated (518A2, 607B, 94.07, A375, 93.08) *TERT*-expressing cell lines subsets regarding chromatin accessibility (two-tailed unpaired t-test; t = 4.63, df = 6; p<0.005). F. Accessibility of mutant allele (%) in a subset of 4 *TERT*p-mutated cutaneous cell lines (518A2, 94.07, A375 and 93.08) calculated as described in Material and Methods (mean ± SD, multiple t-tests, one t-test per cell line, ns p = 0.171; *p<0.001) and the *TERT* mRNA expression in the respective cell lines (mean ± SEM).

In addition, we investigated whether the TERT accessibility originated from the mutant or the wild-type allele. For this purpose, we assessed the fractional abundance of mutated allele, in the subgroup of 4 TERTp-mutated cutaneous cell lines before and after nuclease digestion. 607B cell line was not included since it is homozygous for the mutation and not informative. In 3 out of 4 cell lines preferential digestion of the mutant allele showed that mutated alleles were more accessible than WT alleles (Fig 6F).

Discussion

By using advanced quantitative methods, we investigated the epigenetic and genetic regulation of TERTp in benign and malignant skin cells. Innovative ddPCR-based assays were developed and validated to assess TERT promoter methylation and chromatin accessibility. These methods avoid semi-quantitative qPCR and provide absolute quantification even in samples that are challenged by CG-rich DNA sequences, low concentration and integrity.

In the present study the methylation fraction was assessed by NGS interrogating 31 CpGs in the TERTp region across 44 healthy, benign and malignant tumour samples. Remarkably, high methylation levels were observed in a variety of normal samples. Mainly in keratinocytes methylation levels exceeded those of cutaneous melanoma cell lines. Previous studies on brain tumours and skin melanoma, observed a general absence of methylation in a specific CpG in TERTp, cg11625005, in healthy control samples [3, 20]. Of note, although the authors state absence of methylation we can observe a β-value of ~0.4 (fluorescence ratio provided by Illumina 450K array, ranging from 0 to 1) in their normal samples [3]. In our cohort, the methylation fraction at this CpG was quantified by ddPCR, which validated our results obtained through NGS. Moreover, in our study, methylation of cg11625005 did not stand out across the CpGs in TERTp but seemed to be affected along with adjacent CpGs in this genomic region in all samples (Fig 7A). This result suggests that context-related methylation around cg11625005 is biologically relevant as opposed to methylation of one specific CpG.

TERTp mutations has been described as a genetic mechanism responsible for induction of TERT reactivation [7, 9]. Over the years that followed, a variety of epigenetic or genetic alterations in the gene body or TERTp have been identified, such as promoter methylation, mutations, structural variations, DNA amplification, or promoter rearrangements [3, 5, 19].

In accordance with previous studies, regardless of the methylation status, human benign cells neither harbour TERTp mutation nor express TERT, thereby supporting the principal oncological concept that a benign cell does not undergo undefined proliferation (Fig 8).

Fig 8 — Regardless of MF at the *TERT*p methylation region, both keratinocytes and melanocytes do not show TERT expression. In *TERT*p-mutated cell lines, an intermediate MF positively correlated with chromatin accessibility, in combination with C228T/C250T *TERT* mutations allows monoallelic *TERT* expression. In *TERT*p-WT cell lines, the MF is close to 100% with a significantly higher chromatin accessibility leading to high expression levels. Chromatin schemes adapted from Schwartz *et al*. [26].

Although we have not found a positive correlation between presence of TERTp mutations or TERTp methylation levels and mRNA expression values, all tumour cell lines showed TERT expression, supporting that these mechanisms contribute to telomerase-activation in cancer, separate or in combination [7, 9, 19].

A plethora of histone modifications result in chromatin remodelling that may change accessibility of the TERTp to transcription factors, such as ETS/TCF [7]. Schwartz et al. state that the degree of chromatin folding is correlated with gene transcription and is thought to impact the regulation of DNA-dependent processes [26]. Therefore, we explored the level of chromatin accessibility and its interaction with methylation levels and mRNA expression in 6 cutaneous and 2 uveal melanoma cell lines. In fact, we found a positive correlation between chromatin accessibility and methylation levels as well as mRNA expression that ultimately explains the correlation between methylation fraction and TERT expression. Then, we investigated whether both wild-type and mutant alleles were equally affected by similar patterns of chromatin organization and assessed the mutational fraction upon digestion with nuclease in heterozygous cell lines, assuming that the nuclease only digests DNA open chromatin regions. We could infer that, mutated alleles are more accessible, possibly favouring the binding of transcription factors and consequently TERT mono-allelic expression. Our findings in the 518A2 cell line, harbouring the C228T TERTp mutation, are similar to the results from a study by Stern et al., in which it was found that the active mutant allele is hypomethylated [27]. These observations are consistent with the canonical influence of methylation on transcriptional regulation. In contrast, 94.07 cell line also presents a very small methylation fraction. However, both alleles were equally resistant to nuclease digestion, which might explain the lowest TERT expression levels among all cell lines. Therefore, it still supports the link between local chromatin accessibility and gene regulation [26]. To fully disclose the molecular mechanisms behind TERT expression the heterozygous mutant cell lines A375 and 93.08 provide good models as they allow to study a repressed and expressed allele within the same cell.

Another remarkable observation in our study is that in WT TERT-expressing uveal melanoma cell lines, the methylation of the whole TERTp region is close to 100% with a significantly higher chromatin accessibility compared to TERTp-mutated cell lines. Accordingly, Stern et al. also demonstrate that cell lines with WT TERTp display much higher levels of methylation [27]. These characteristics of WT TERTp cell lines may lead to biallelically TERT activation under distinct epigenetic conditions from those in mutated TERTp.

Interestingly, these results suggest a complex model in which TERT expression requires either a widely open chromatin state in TERTp-WT samples due to hypermethylation throughout the promoter or mono-allelic expression of the accessible mutated allele in combination with moderate (probably allele-specific) methylation fraction (Fig 8). Furthermore, Huang and colleagues reported that some cancer cell lines show mono-allelic expression of TERT even in the absence of TERTp mutations [28].

Previous studies have reported the association between TERTp hypermethylation and poor patient survival in melanoma and other cancers, indicating that it might be a relevant prognostic marker [20, 27, 29–31]. In primary melanoma it needs to be assessed if TERTp methylation is predictive of worse prognosis. Thus, the quantification of TERT methylation through ddPCR might be relevant in the clinic to assess patient prognosis.

The dynamics of epigenetic mechanisms in TERT genetic regulation is complex. Further investigations are needed to address the correlation of allele-specific differences in chromatin accessibility and promoter methylation with allele-specific mRNA expression.

Material and methods

Samples, DNA extraction and PCR

Surplus female breast skin and nevi tissues were obtained from 11 and 6 anonymous patients that underwent cosmetic surgery, respectively. Surgeries for mama reduction (performed between 2010 and 2018) and naevi (performed between 2008 and 2009), were conducted according to declaration of Helsinki principles. Epidermis and dermis were separated after removal of adipose tissue followed by enzymatic digestion and primary fibroblast (n = 5) and keratinocyte (n = 8) cell suspensions were obtained and cultured as described before [32]. Keratinocytes were used at passage 2, while fibroblasts were used at passage 3–5.

Low-passage cultured melanocytes (n = 5)–m003, m003A, m002, m004A and 0398A –were cultured as previously described [33]. HEMs were cultured more recently in the medium 254 supplemented with HMGS-2 (Gibco/ThermoFisher) and Penicillin (100 U/ml), and Streptomycin (100 μg/ml; both from Lonza, Verviers, Belgium).

We also included 19 early-passage cutaneous melanoma cell lines derived from metastatic lesions cultured for research purposes and adoptive T-cell transfer [34]. Cell lines were cultured and DNA and RNA extracted between 2017 and 2019. The 518A2, 607B, 04.01, 04.04, 94.13, 93.05, 94.07, 93.08, 634, 01.05, and 06.24 cell lines were a kind gift from Dr. Els Verdegaal (Department of Medical Oncology, LUMC). Meljuso was obtained from Prof. Neefjes (Department of Cell and Chemical Biology, LUMC). WM1361A, WM3506, WM1960 cell lines were a kind gift from Dr. KL Scott (Baylor College of Medicine, Houston, USA). MM057 and A375 were kindly provided by Prof. JC Marine (VIB, Leuven, Belgium). OCM8 and OCM1 were provided by Dr. Mieke Versluis (Department of Ophthalmology, LUMC) [35]. All cell lines were cultured with Dulbecco’s modified eagle medium (DMEM, low glucose, pyruvate; Gibco/ThermoFisher) supplemented with 10% FCS, Penicillin (100 U/ml), and Streptomycin (100 μg/ml; both from Lonza, Verviers, Belgium) and glutamax (100X, Gibco).

For the 6 uveal cell lines provided by Dr. Mieke Versluis (Department of Ophthalmology, LUMC), the establishment and culturing conditions have been described before: OMM 1 [36], OMM 2.3, OMM 2.5 and Mel270 [37], Mel202 [38], 92.1 [39]. All cell lines used in our study were tested negative for mycoplasm and recently subjected to STR profiling.

The batch thus consisted of 36 primary skin type samples and 25 melanoma cell lines, totalling 61 samples (Table 1).

Table 1. Samples overview.

Control samples					Melanoma cell lines
Skin biopsy samples	Fibroblasts	Melanocytes	Keratinocytes	Naevi	Cutaneous	Uveal
LB627	F537	m003	K590	Naevus 1	04.01	OMM 2.3
LB470	F544	m002	K409	Naevus 2	WM1361A	OMM 1
LB579	F332	m003A	K549	Naevus 3	93.05	OMM 2.5
LB576	F334	m004A	K514	Naevus 4	WM3506	Mel270
LB584	F628	0398A	K060	Naevus 5	WM1960	Mel202
LB586		HEM	K627	Naevus 6	Meljuso	92.1
LB625			K516		634
LB381			K550		OCM8
LB628					OCM1
LB629					518A2
Fresh skin 1					607B
					94.07
					A375
					93.08
					94.13
					01.05
					04.04
					MM057
					06.24

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DNA was isolated using the QIAamp DNA Blood Mini Kit and the DNeasy Blood & Tissue Kit (both from Qiagen, Hilden, Germany).

Conventional PCR was performed using the PCR-sequencing kit (Thermo Fisher Scientific, Waltham, MA, USA), containing 10X reaction buffer, MgCl2 (50mM), dNTP mix (10nM, Fermentas/Thermo Fisher Scientific), primer mix (900nM each), PlatinumX Taq enzyme (2.5U), 50ng DNA and Aqua B. Braun RNase-free water. A PCR for CG-rich sequences was performed on 50ng DNA using the PCRX Enhancer System (Thermo Fisher Scientific), containing 10X PCRX amplification buffer, MgSO4 (50mM), dNTP mix (10nM), primer mix (900nM each), PlatinumX Taq enzyme (2.5U) and Aqua B. Braun RNase-free water. The samples were amplified in C1000 Touch Thermal Cycler (Bio-Rad Laboratories, Inc., Hercules, CA, USA).

Promoter methylation determination

Bisulfite conversion and next-generation sequencing (NGS)-based deep bisulfite sequencing

In this experiment 44 samples were included: fibroblasts (n = 5), melanocytes (n = 5), naevi (n = 6), normal skin samples (n = 11), keratinocytes (n = 8), cutaneous melanoma cell lines (n = 6) and uveal melanoma cell lines (n = 3). DNA was bisulfite-converted (BC) using the EZ DNA Methylation^™ Kit (Zymo Research, Irvine, CA, USA) according to the manufacturer protocol (version 1.2.2). BC samples were amplified using the PCRX Enhancer System in the program: 1 cycle of 95°C for 3 minutes, 8 cycles of 95°C for 30 seconds, 58°C for 30 seconds, reducing 1°C/cycle, and 68°C for 1 minute, then 36 cycles of 95°C and 53°C for 30 seconds each, and 68°C for 1 minute, followed by 1 cycle of 68°C for 3 minutes. Tailed primers were used for amplification (900nM each; S1 Table). Samples were sequenced through next-generation sequencing (NGS), MiSeq, 2x300bp paired-end, at Leiden Genome Technology Centre (LGTC). Bisulfite sequencing reads were quality trimmed using PRINSEQ (v0.20.4 lite) and aligned to GRCh37 using Bismark (v0.20.0) and Bowtie 2 (v2.3.4.3) [40–42].

Novel design of a ddPCR assay using methylation-sensitive restriction enzymes (MSREs) to determine TERTp methylation fraction

The methylation fraction (MF) of the CpG (cg11625005) in position 1,295,737 was determined by an in-house designed ddPCR assay in combination with HgaI methylation-sensitive restriction enzyme (MSRE) that cleaves this CpG when unmethylated, as described by Nell et al. [24]. 100ng DNA sample was incubated with HgaI (2U/μl) and appurtenant 10X NEBuffer 1.1 (both from New England Biolabs, Bioké, Leiden, The Netherlands) for 60 minutes at 37°C and 65°C for 20 minutes. To assess the MF of a CpG adjacent to cg11625005, located in 1,295,731, the MSRE AvaI (10U/μl; New England Biolabs) was employed, which recognises this CpG and cleaves it when unmethylated. Incubation of the DNA samples with AvaI was performed with 10X CutSmart buffer for 15 minutes at 37°C and subsequently 65°C for 20 minutes. For ddPCR reaction, 60ng DNA digested or undigested by HgaI, 2x ddPCR SuperMix for Probes (no dUTP), primers (900nM each), a FAM-labelled in-house-designed probe for the CpG site of interest (250nM, Sigma, St. Louis, MO, USA), and 20X HEX-labelled CNV TERT reference primer/probe (Bio-Rad) for total TERT amplicon count. The primers and probe sequences are presented in S2 Table. The amplification protocol used: 1 cycle of 95°C for 10 minutes, 40 cycles of 94°C for 30 seconds and 60°C for 1 minutes, and 1 cycle of 98°C for 10 minutes, all at ramp rate 2°C/s. Droplets were analysed through a QX200 droplet reader (Bio-Rad) using QuantaSoft software version 1.7.4 (Bio-Rad). Raw data was uploaded in online digital PCR management and analysis application Roodcom WebAnalysis (version 1.9.4, https://www.roodcom.nl/webanalysis/) [24], in which the MF was calculated by dividing the CNV of the digested sample with that of the paired undigested sample.

Assessment of mutational status

Sanger sequencing

The presence of the C228T and C250T TERTp mutations in some samples was evaluated by conventional Sanger sequencing. DNA samples were amplified through the PCRX Enhancer System (Thermo Fisher Scientific) using primers (Sigma-Aldrich) and amplification program described by McEvoy et al. [25].

Mutation analysis using commercial TERT C250T and C228T mutation assays

For most of the samples, the TERTp mutations were detected by the ddPCR technique according to protocol described by Corless et al. [43], using the TERT C250T_113 Assay and C228T_113 Assay (unique assay ID dHsaEXD46675715 and dHsaEXD72405942, respectively; Bio-Rad). Both assays include FAM-labelled probes for the C250T and C228T mutations respectively, HEX-labelled wild-type (WT) probes, and primers for a 113-bp amplicon that encompasses the mutational sites. The ddPCR reaction mix comprised 1X ddPCR Supermix for Probes (No dUTP), Betaine (0.5M; 5M stock), EDTA (80mM; 0.5M stock, pH 8.0, Thermo Fisher Scientific), CviQI restriction enzyme (RE; 2.5U; 10U/μl stock, New England BioLabs), the TERT assay, and 50ng DNA. Droplets were generated in QX200 AutoDG system (Bio-Rad) and amplified in T100 Thermal Cycler (Bio-Rad) according to the recommended cycling conditions and analysed through a QX200 droplet reader (Bio-Rad) using QuantaSoft software version 1.7.4 (Bio-Rad).

Chromatin accessibility

Cell culture and treatment to assess chromatin states

Cutaneous melanoma cell lines A375, 518A2, 607B, 94.07, 93.08, OMM2.5, Mel270 and OCM8 were cultured for 22 days in 9-cm Cellstar^® cell culture dishes (Greiner Bio-One GmbH, Frickenhausen, Germany) with Dulbecco’s modified eagle medium (DMEM; Sigma-Aldrich) supplemented with 10% FCS, Penicillin (100U/ml), and Streptomycin (100μg/ml; both from Lonza, Verviers, Belgium) until roughly 95% confluent. Then, different densities (10,000, 20,000, 40,000 and 80,000 cells) of the above-mentioned cell lines were seeded in duplicate into a 48-well plate (Corning Costar, Sigma-Aldrich) required for the EpiQ chromatin assay. The EpiQ^™ Chromatin Analysis Kit (Bio-Rad) was performed according to manufacturer’s instructions. Briefly, after 2 days each cell line was 85%-95% confluent. The cells were permeabilised and treated with EpiQ chromatin digestion buffer with or without nuclease for 1 hour at 37°C. Following incubation with EpiQ stop buffer for 10 minutes at 37°C, the DNA samples were purified using alcohol and DNA low- and high-stringency wash solutions. The genomic DNA was eluted in DNA elution solution.

Novel design of a ddPCR assay to assess chromatin opening state

The analysis was performed using ddPCR rather than qPCR, to achieve quantifiable results using GAPDH expression as positive control. The reaction mix consisted of 2x ddPCR Supermix for Probes (No dUTP, Bio-Rad), 20x HEX-labelled CNV TERT reference primer/probe (Bio-Rad), 50ng DNA, and primers (900nM each) and FAM-labelled probes (250nM) for GAPDH, or the methylation region around cg11625005 (S3 Table). Samples were amplified according to the program of the CNV TERT reference primer/probe as described. Gene accessibility was quantified by the digestion fraction between the digested and undigested samples, subtracted from 1, multiplied by 100.

Allele-specific chromatin accessibility

The mutational fraction upon digestion with nuclease (EpiQ^™ Chromatin Analysis Kit aforementioned) was assessed in cutaneous melanoma cell lines with heterozygous TERTp mutations, 518A2, 94.07, A375 and 93.08. The analysis was performed by ddPCR using the TERT C250T_113 Assay and C228T_113 Assay (unique assay ID dHsaEXD46675715 and dHsaEXD72405942, respectively; Bio-Rad) as described above. The mutation fraction from undigested and digested samples were compared and the accessibility of mutant allele was calculated as follows:

A c c e s s i b i l i t y o f m u t a n t a l l e l e = \frac{m u t a t i o n a l f r a c t i o n u n d i g e s t e d - m u t a t i o n a l f r a c t i o n d i g e s t e d}{m u t a t i o n a l f r a c t i o n u n d i g e s t e d} \times 100

RNA isolation, cDNA synthesis and quantitative real-time PCR

RNA was obtained using the FavorPrep Tissue Total RNA Extraction Mini Kit (Favorgen Biotech, Vienna, Austria) according to manufacturer’s instructions for animal cells. cDNA was synthesised through the iScript^™ cDNA Synthesis Kit (Bio-Rad) according to recommended protocol. TERT mRNA expression was assessed by qPCR performed with 3.5ng DNA, IQ SYBR Green Supermix (2x; Bio-Rad), and 0.5μM PCR primers (Sigma-Aldrich; S4 Table) in a Real-Time PCR Detection System CFX96 (Bio-Rad) and normalised to reference gene expression (RPS11, TBP and CPSF6, S4 Table). Data was analysed through the ΔΔCT method in Bio-Rad CFX manager software (version 3.1, Bio-Rad).

Statistical analysis

In this study we used the GraphPad Prism software (version 8.0.1 for Windows, GraphPad Software, CA, USA) to perform all the statistical tests. Prism 8 has a wide library of analysis and in our paper we have used the linear regressions and correlations (in Figs 2A, 2C, 2D, 3; 6B, 6C, 6D), one-way ANOVA (Fig 5) and multiple t-tests without correction for multiple comparisons, one t-test per cell line, *p<0.001). (Fig 6A and 6F) and two-tailed unpaired t-test (Fig 6E). A p-value<0.05 was considered statistically significant.

The methylation fraction obtained using ddPCR was calculated with 95% confidence interval by dividing the CNV of the digested sample with that of the paired undigested sample. Raw data was uploaded in online digital PCR management and analysis application Roodcom WebAnalysis (version 1.9.4, https://www.roodcom.nl/webanalysis/) [24] (in Fig 2B).

Ethics statement

The study was conducted according to the Declaration of Helsinki Principles.

Naevi samples from 6 patients (excised between 2008 and 2009) were accessed anonymously from the biobank of the Department of Dermatology, LUMC and was approved by the Leiden University Medical Center institutional ethical committee (05–036).

Surplus female breast skin was obtained from 11 anonymous patients that underwent surgeries for mama reduction (performed between 2010 and 2018). This entails patient consent was not required since the surplus tissue was considered as waste material. Experiments were conducted in accordance with article 7:467 of the Dutch Law on Medical Treatment Agreement and the Code for proper Use of Human Tissue of the Dutch Federation of Biomedical Scientific Societies (https://www.federa.org/codes-conduct). As of this national legislation, coded tissue samples can be used for scientific research purposes when no written objection is made by the informed donor. Therefore, additional approval of an ethics committee regarding scientific use of surplus tissue was not required.

Supporting information

S1 Table. Tailed primers used for amplification of 325-bp region in bisulfite-converted samples.

(XLSX)

Click here for additional data file.^{(9KB, xlsx)}

S2 Table. Primers and probe sequences to amplify the 106-bp amplicon in a novel design of a ddPCR assay to determine the methylation fraction.

(XLSX)

Click here for additional data file.^{(9KB, xlsx)}

S3 Table. Primers and probe sequences to amplify the 231-bp region encompassing 31 CpG sites around the cg11625005 in a novel ddPCR assay to assess the chromatin state.

(XLSX)

Click here for additional data file.^{(9.1KB, xlsx)}

S4 Table. Primer and probe sequences for TERT expression in qPCR.

(XLSX)

Click here for additional data file.^{(9.2KB, xlsx)}

S5 Table. Overview of the methylation fraction (measured by ddPCR and NGS), mutational status and TERT mRNA expression of our sample cohort (n = 61).

(XLSX)

Click here for additional data file.^{(17.5KB, xlsx)}

S6 Table. Overview of the methylation fraction (measured by ddPCR and NGS), mutational status and TERT mRNA expression and chromatin accessibility in the subset of melanoma cell lines present of our cohort (n = 25).

(XLSX)

Click here for additional data file.^{(14.4KB, xlsx)}

S7 Table. Raw data used in the Fig 5.

(XLSX)

Click here for additional data file.^{(9.2KB, xlsx)}

S8 Table. Raw data used in Fig 6.

(XLSX)

Click here for additional data file.^{(17.2KB, xlsx)}

S9 Table. Results overview.

(XLSX)

Click here for additional data file.^{(33.1KB, xlsx)}

Acknowledgments

We thank Wim Zoutman, Abdoel el Ghalbzouri, AG Jochemsen and Mijke Visser for useful discussions, Marion Rietveld, Coby Out and Tim van Groningen for the assistance with cell culturing. We would like to thank Dr. Mieke Versluis (Department of Ophthalmology, LUMC), Dr. Els Verdegaal (Department of Medical Oncology, LUMC), Prof. Neefjes (Department of Cell and Chemical Biology, LUMC), Dr. KL Scott (Baylor College of Medicine, Houston, USA) and Prof. JC Marine (VIB, Leuven, Belgium) for kindly providing melanoma cell lines.

Data Availability

All relevant data are within the manuscript and its Supporting Information file.

Funding Statement

This project has received funding from the European Union’s Horizon 2020 research and innovation programme under the grant agreement No 641458 (http://melgen.org/). RN is supported by the European Union’s Horizon 2020 research and innovation program under grant agreement No 667787 (UM Cure 2020 project, https://www.umcure2020.org/en/). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

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PLoS One. doi: 10.1371/journal.pone.0231418.r001

Decision Letter 0

Srinivas Saladi

Transfer Alert

This paper was transferred from another journal. As a result, its full editorial history (including decision letters, peer reviews and author responses) may not be present.

31 Dec 2019

PONE-D-19-33667

Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression

PLOS ONE

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Reviewer #1: Summary:

The authors study genetic and epigenetic contribution to TERT gene reactivation in the context of normal skin cells and melanomas. They analyzed 44 primary tissue samples and cell-lines and quantified the methylation fraction (NGS based deep bisulphite sequencing and ddPCR assay) and mutational status (sanger sequencing and ddPCR assay) of the TERT gene. Interestingly, though the authors found that previously characterized mutations in TERT gene were unique to melanoma cell-lines, they did not seem to be correlated to expression. Indeed TERT expression seemed to be exclusive to melanoma cell-lines regardless of mutation status. Similarly, the methylation fraction of a cell-type also did not seem to be correlated to TERT expression levels. Both these findings are contrary to what has been previously characterized about TERT. In an effort to explain this discrepancy, the authors also investigated chromatin accessibility. They found that accessibility around a specific CpG was correlated to methylation and mRNA expression levels in the 8 cell-lines investigated.

Major Comments:

The study is well designed and done in both primary tissue samples and cell-lines. It furthers our understanding of the complexity of TERT gene and hence is of significance to the field.

Minor Comments:

Some things that can be done to make the manuscript more impactful:

1. The authors findings are in direct contradiction to previously carried out studies on TERT. Their accessibility experiments add another layer of complexity but do not explain the discrepancy in the relationship between methylation/mutation status and expression. It would be helpful if the authors postulate the reasons for these contradictions and discuss the impact of these findings in greater detail.

2. Figures submitted are of low resolution, as such it was hard to verify cell-line names, legends etc.

3. In Methods Statistical Analysis: mention the exact libraries used in the softwares so that the analysis can be replicated

4. Provide the raw tables that were used to make the box plots and other graphs

Reviewer #2: Salgado et al report on the association between TERT promoter mutations, DNA methylation, and chromatin accessibility in melanoma and normal cells. The authors characterize methylation fractions using bisulfite sequencing of normal skin as well as different cell types, naevi, and melanoma cell lines, finding that uveal melanoma cell lines have the highest levels of methylation (Fig. 2). Using ddPCR, they then determine that keratinocytes have a higher methylation level at a position on the TERT promoter than melanoma cell lines (Fig. 2). There was no correlation between TERT methylation at a previously characterized position and mRNA expression when comparing 34 samples and cells (Fig. 3). However, while TERT promoter mutations were identified in some melanoma cell lines and tissue (Fig. 4), there was no correlation with mRNA expression (Fig. 5). Since TERT expression was only seen in melanoma cells lines, additional analysis was conducted on these. It was then found that there was a high degree of variability in chromatin accessibility at cg11625005 between different melanoma cell lines as well as a correlation with the methylation fraction (Fig. 6). In a subset of 8 melanoma cell lines, there is a correlation between MF, chromatin accessibility, and mRNA expression of TERT. Allele specific analysis of chromatin accessibility indicated that the mutated alleles were more accessible than wildtype.

This study nicely shows that ddPCR can generate high quality methylation data that validates bisulfite sequencing. However, it is not clear that the results of this study are a big enough advance for publication. The findings are descriptive rather than mechanistic and often unsubstantiated. The allele specific data regarding chromatin accessibility are interesting but should be expanded. The correlation between high levels of chromatin accessibility at the TERT promoter and mRNA levels is also interesting but not a mechanistic finding on its own. It would be more informative to elucidate a mechanism for the varying levels of chromatin accessibility like determining transcription factor or chromatin remodeling enzyme association. Also, the nuclease assay used in this study likely determines chromatin structure at the nucleosome level and not higher order chromatin structure.

Specific points:

1. Overall, the conclusions of this study are not very clear because of the presentation of conflicting pieces of data. The lack of correlations in tissues conflicts with the correlations found in cell lines. The discussion should do a better job in explaining the physiological significance of the latter studies on a subset of melanoma cell lines.

2. It is interesting that there is allele specific differences in chromatin accessibility. Does this correlate with allele specific mRNA expression? Is there a correlation between ETS/TCF binding and chromatin accessibility at the mutated allele? It is possible that chromatin immunoprecipitations to investigate ETS binding may be informative. Otherwise, the authors should investigate an alternative hypothesis for investigating a mechanism.

3. The graphs in Fig. 6A should include error bars and both Fig.6A and Fig.6F should include p values.

4. There should be more information on the different assays used, especially the chromatin accessibility assay. Is this assay based on nuclease digestion of permeablized nuclei? If so, then it does not measure higher order chromatin structure.

5. Figure legends should include more information about the tissues and cell lines used for the specific study.

6. Lines 85 to 89 in the Results section do not describe the results in Fig. 1 very well. The authors should call attention to the finding that melanocytes are among the lowest. The difference between normal melanocytes and cutaneous melanoma cells is more than comparisons between melanomas and keratinocytes or fibroblasts.

7. The text should be modified to avoid usage of “higher order chromatin structure”.

**********

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PLoS One. 2020 Apr 8;15(4):e0231418. doi: 10.1371/journal.pone.0231418.r002

Author response to Decision Letter 0

18 Feb 2020

Dear Editor,

First we would like to express our appreciation for considering our manuscript entitled “Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression” for publication in PLOS ONE and for the thoughtful reviews provided by the referees. We have attempted to address all comments and adapted the manuscript accordingly. Please find our responses below:

Reviewer #1:

Major Comments:

The study is well designed and done in both primary tissue samples and cell-lines. It furthers our understanding of the complexity of TERT gene and hence is of significance to the field.

Answer:

We thank the reviewer for the positive and constructive comments.

Minor Comments:

Some things that can be done to make the manuscript more impactful:

Answer:

In our study we explore a big cohort encompassing both control/benign samples and melanoma cell lines.

Previous studies stated absence of methylation in the TERTp in benign samples, while we found high methylation fractions (MF) in some healthy skin samples, such as normal skin (~30%), naevi (~30%) and cultured keratinocytes (~50%). In fact, in keratinocytes, the MF was as high as in cutaneous melanoma cell lines. We found this result quite intriguing, but we think it can be explained by technical differences and differences in interpretation. Whereas we used a validated quantitative approach previous studies use semi-quantitative approaches in which they interpreted a β-value of ~0.4 in normal tissue as not methylated. Therefore, the measurements seem to be similar to previous studies and it is rather a difference in interpretation. Moreover, in accordance to previous studies none of the human-derived benign samples neither harbour TERTp mutation nor expressed TERT, thereby supporting the basic oncological concept that a benign cell does not undergo undefined proliferation.

Analysis of tumour cell lines revealed a wide range of promoter methylation levels (from 5% to 100%). Besides promoter methylation, somatic mutations are also known to be correlated with TERT reactivation. Therefore, we characterized the TERTp mutational status of the sample cohort. While 12 out of 25 showed C250T mutation and 5 the C228T mutations, 8 out of 25 were WT. All melanoma cell lines showed TERT expression, (n=25) irrespective of mutation and methylation status, supporting that there are additional telomerase-activating mechanisms involved in cancer besides methylation and mutations in TERTp [1-3]. This holistic model in which mutation, methylation of TERTp and other unidentified mechanisms might explain TERT expression in WT melanoma cell lines.

Schwartz et al. state that the degree of chromatin folding is correlated with gene transcription and is thought to impact the regulation of DNA-dependent processes [4]. Other studies demonstrated disease-dependent differences in nucleosome positions in tissues and cell lines, suggesting an important role of chromatin architecture on cell fate [5]. Moreover, chromatin accessibility is modulated locally, since nuclease-accessible sites were enriched at gene regulatory elements, such as active TSS, indicating a link between local chromatin accessibility and gene regulation [4]. TERTp is reported to coincide with a DNAse 1 hypersensitive locus as evidenced in publicly available data (UCSC database, DHS track) which supports that accessibility at this site is involved in gene regulation.

Since the accessibility experiments require cell culturing we were not able to perform it in the benign cells since the cultured keratinocytes, fibroblasts and melanocytes are isolated at low passages. Thus, we explored the level of chromatin accessibility and TERT expression in 8 cell lines with different methylation/mutation patterns. In fact, we found a positive correlation between chromatin accessibility and methylation levels as well as mRNA expression that ultimately explains the correlation between methylation fraction and TERT expression. In WT TERT-expressing uveal melanoma cell lines, the methylation of the whole TERTp region is close to 100% with a significantly higher chromatin accessibility compared to TERTp-mutated cell lines. Our results also show that mutated alleles in heterozygous cell lines may be more accessible and allows further dissection of the underlying mechanisms that facilitate mono-allelic expression.

Interestingly, these results suggest a complex model in which TERT expression requires either a widely open chromatin state in TERTp-WT samples due to hypermethylation throughout the promoter leading to biallelically TERT activation or mono-allelic expression of the accessible mutated allele in combination with moderate (probably allele-specific) methylation fraction. Most of these results are in agreement with the findings from Stern et al. [6].

This model supported our view that the integration of different aspects are definitely more important than studying each one separately.

To further address this comment we have clarified the assumptions underlying this study and refined the conclusions in the manuscript text.

2. Figures submitted are of low resolution, as such it was hard to verify cell-line names, legends etc.

Answer:

We have verified the figures and adapted the figure resolution, the font size of cell lines and legends in agreement with journal requirements.

3. In Methods Statistical Analysis: mention the exact libraries used in the softwares so that the analysis can be replicated.

Answer:

In this study we used the GraphPad Prism software (version 8.0.1 for Windows, GraphPad Software, CA, USA) to perform all the statistical tests. Prism 8 has a wide library of analysis and in our paper we have used the linear regressions and correlations, one-way ANOVA, multiple t-tests (one t-test per cell line) and two-tailed unpaired t-test. A p-value<0.05 was considered statistically significant.

A detailed statistical analysis subsection meeting the journal requirements has been added to the material and methods section of the manuscript.

4. Provide the raw tables that were used to make the box plots and other graphs

Answer:

To address this point a supplementary excel file has been created containing the raw data behind each plot in different sheets.

Reviewer #2:

Salgado et al report on the association between TERT promoter mutations, DNA methylation, and chromatin accessibility in melanoma and normal cells. The authors characterize methylation fractions using bisulfite sequencing of normal skin as well as different cell types, naevi, and melanoma cell lines, finding that uveal melanoma cell lines have the highest levels of methylation (Fig. 2). Using ddPCR, they then determine that keratinocytes have a higher methylation level at a position on the TERT promoter than melanoma cell lines (Fig. 2). There was no correlation between TERT methylation at a previously characterized position and mRNA expression when comparing 34 samples and cells (Fig. 3). However, while TERT promoter mutations were identified in some melanoma cell lines and tissue (Fig. 4), there was no correlation with mRNA expression (Fig. 5). Since TERT expression was only seen in melanoma cells lines, additional analysis was conducted on these. It was then found that there was a high degree of variability in chromatin accessibility at cg11625005 between different melanoma cell lines as well as a correlation with the methylation fraction (Fig. 6). In a subset of 8 melanoma cell lines, there is a correlation between MF, chromatin accessibility, and mRNA expression of TERT. Allele specific analysis of chromatin accessibility indicated that the mutated alleles were more accessible than wildtype.

study nicely shows that ddPCR can generate high quality methylation data that validates bisulfite sequencing. However, it is not clear that the results of this study are a big enough advance for publication. The findings are descriptive rather than mechanistic and often unsubstantiated. The allele specific data regarding chromatin accessibility are interesting but should be expanded. The correlation between high levels of chromatin accessibility at the TERT promoter and mRNA levels is also interesting but not a mechanistic finding on its own. It would be more informative to elucidate a mechanism for the varying levels of chromatin accessibility like determining transcription factor or chromatin remodeling enzyme association. Also, the nuclease assay used in this study likely determines chromatin structure at the nucleosome level and not higher order chromatin structure.

Specific points:

Answer:

Throughout the last 20 years many processes have been identified as having an effect in the regulation of TERT gene, namely histone modifications, such as acetylation, methylation, phosphorylation, and ubiquitinization [8], CpG methylation at TERTp and TERTp mutations. TERT expression was associated with hyperacetylation of core histones at the TERTp [9, 10]. Methylation at lysine 9 of histone H3 (H3K9) and lysine 20 of H4 (H4K20) were features of telomerase-negative immortal cells, whereas methylation at lysine 4 of histone H3 (H3K4) was usual in telomerase-positive cells [11]. Depletion of the histone methyltransferase SMYD3 leading to reduction of histone H3K4 methylation at TERTp led to downregulation of TERT [12] and inhibition of demethylase LSD1 led to increase of methylation at H3K4 and an TERT upregulation [13].

A CpG island covering the TERTp has caught attention of the scientific community since in cancer cells and cell lines [14, 15] it was often methylated but the correlation with transcription has been hard to make due to the CG-rich sequence. It has been suggested that hypermethylation was implicated in the positive regulation of the TERTp [16] possibly due to the hampering of binding of transcriptional repressors [17].

Only few years ago with the advent of NGS-technologies, namely NGS-based bisulfite sequencing and nowadays the emerging techniques, as ddPCR that we present in our study, it became possible to draw robust conclusions on how TERTp methylation affects TERT gene regulation [6].

Within the last decade with the discovery of TERTp mutations and the correlation with TERT upregulation, due to the generation of new transcription factors binding motifs, another layer of complexity arose and allowed to clarify the mechanism in some types of cancer [2, 3].

Zhu and colleagues postulated that the repressive chromatin environment and nucleosomal conformation appear to be one of the major mechanisms that tightly suppress the TERT gene in majority of human somatic cells [8].

We can see that all the aforementioned studies have contributed at multiple levels of gene regulation to reveal the complex process of regulating TERT expression.

In our study, we were able to develop and validate innovative ddPCR-based assays to assess TERT promoter methylation and chromatin accessibility in a quantitative fashion.

While previous studies stated absence of methylation in the TERTp in benign samples despite β-values of ~0.4 in methylation array analysis, we presented significant methylation fractions (MF) in some healthy skin samples, such as normal skin, naevi and keratinocytes. However, in accordance to previous studies none of the human-derived benign samples neither harbour TERTp mutation nor expressed TERT, thereby supporting the basic oncological concept that a benign cell does not undergo undefined proliferation.

Analysis of tumour cell lines revealed a wide range of promoter methylation levels (from 5% to 100%) and different TERTp mutational status: 12 out of 25 showed C250T mutation, 5 the C228T mutations and 8 out of 25 were WT. However, all tumour cell lines showed TERT expression, (n=25) irrespective of mutation and methylation status, supporting that there are other telomerase-activating mechanisms in cancer besides methylation and mutation [1-3].

Then, we comprehensively analysed the chromatin status in the light of current knowledge. Back in 2004, Wang and Zhu have described the chromatin environment to be critical for the tight regulation of the TERT gene, stating that in telomerase-positive cells, the TERT transcription was accompanied by the appearance of a major DNase I hypersensitive site (DHS) at the core promoter [18, 19].

In the present study, from the chromatin accessibility assay, we could observe an interplay between DNA methylation and presence of TERTp mutations culminates in different levels of accessibility and thus TERT expression. Our results also suggest that mutated alleles in heterozygous tumors are more accessible compared to WT allele, favoring mono-allelic expression.

These results suggest a complex model in which TERT expression requires either a widely open chromatin state in TERTp-WT samples due to hypermethylation throughout the promoter leading to biallelically TERT activation or mono-allelic expression of the accessible mutated allele in combination with moderate (probably allele-specific) methylation fraction. Most of these results are in agreement with the findings from Stern et al. [6].

We believe that the major interest is to integrate all the data to understand how this important gene involved in immortalization and undefined proliferation is regulated. We hereby provide a holistic model that is based on observations in tissue and cell lines. This offers the possibility to investigate TERT regulation in vitro and in vivo in much higher detail. However, it is beyond of the scope of this article to resolve the actual mechanism.

We have clarified these important points by adapting the discussion section.

Answer:

We thank the reviewer for providing this relevant suggestion. In fact, our results suggest that there are allele specific differences in chromatin accessibility. Showing a correlation with allele specific mRNA expression would be of interest, however the informative cell lines do not provide heterogeneity in the transcribed gene and hence does not allow allele-specific quantification. Differential CTCF binding is definitely a good possibility but is beyond the scope of the present study. However, we state that further investigations are needed to fully disclose this mechanism.

3. The graphs in Fig. 6A should include error bars and both Fig.6A and Fig.6F should include p values.

Answer:

The reviewer’s comment made us see that Figure 6 needs to be adapted. Therefore, in the figure 6A the error bars were added. The GAPDH bars were removed and mentioned in the legend. The TERT methylation region values were normalized and compared to withdrawn GAPDH data in order to generate p-values.

Regarding the Fig 6F, error bars were added in the expression bars. Moreover, we decided to simplify the figure by showing the subtraction between mutational fraction in undigested and digested divided by undigested fraction as follows:

Accessibility of mutant allele =(mutational fraction undigested-mutational fraction digested )/(mutational fraction undigested)×100

Statistically significant differences in mutational fraction before and after nuclease digestion are indicated by asterisks.

Based on this remark we have added an ‘allele-specific chromatin accessibility’ subsection in the material and methods section.

4. There should be more information on the different assays used, especially the chromatin accessibility assay. Is this assay based on nuclease digestion of permeabilized nuclei? If so, then it does not measure higher order chromatin structure.

Answer:

In the material and methods section we state that chromatin analysis was performed according to manufacturer’s instructions followed by a brief description of the method used “The cells were permeabilized and treated with EpiQ chromatin digestion buffer with or without nuclease for 1 hour at 37°C”. In fact, the kit used to evaluate the chromatin status is based on nuclease digestion of permeabilized nuclei. Since it is part of a commercial kit the type of nuclease is not revealed. Like that we could not explore the reported function and better interpret the readout of our experiment, as performed for MNase endonuclease used in the study of Schwartz et al., 2018 [4]. However, since the site that we investigate, overlaps with a DNase 1 hypersensitive site we assume that our test can also detect a repressive chromatin. Moreover, this is supported by a good correlation between gene accessibility and gene expression (Fig. 6C).

Therefore, through this methodology we were able to measure chromatin accessibility that is modulated on local scale, with nuclease-accessible sites enriched at gene regulatory elements, such as active TSS [4]. To accurately measure the higher order chromatin structure we would need to perform a genome-wide chromatin immunoprecipitation (ChIP) analyses to assess the dynamics of higher-order chromatin organization [20] or even more precise a DNA-labelling method to stain the higher-order chromatin clusters and fibers that enables the reconstruction of ultrastructure to be visualized in the nucleus of human cells in situ, such as the ChromEMT (ChromEM tomography) [21].

As above mentioned, the material and methods section has been edited in order to give more detailed information about the assays used, namely about the chromatin accessibility assay for which an ‘allele-specific chromatin accessibility’ subsection has been added.

5. Figure legends should include more information about the tissues and cell lines used for the specific study.

Answer:

The figure legends have been adapted and now they incorporate information about cell lines and tissues used and also some details about statistical analysis performed.

Answer:

We thank the reviewer for this constructive comment. We have stressed the difference in methylation fraction between keratinocytes and melanoma cell lines, since has been reported that there is an absence of methylation at these location in non-malignant samples [22, 23], while we found similar methylation levels in both keratinocytes and melanoma cell lines. However, we agree that since melanocytes are the cellular origin of melanoma it is quite remarkable that we found this striking difference in methylation between these two types of samples.

The explanation of Fig. 1 in the results section has been extended.

7. The text should be modified to avoid usage of “higher order chromatin structure”.

Answer:

We thank the reviewer for drawing attention to this potentially confusing point. Most textbooks present the long-lasting model in which the primary DNA-nucleosome subunits progressively fold into discrete higher-order chromatin fibers and, ultimately, mitotic chromosomes. Despite all the research carried out the chromatin organization is still an enigma. However, recent literature reports that no global differences in DNA packaging do exist, challenging the model of hierarchically organized higher-order structures of chromatin [4, 21].

The manuscript text has been adapted in order to avoid the expression.

We thank the reviewers for their thoughtful comments. Addressing the questions and comments has helped us to clarify parts of the study. We have the impression that the adaptations have improved the manuscript considerably.

Kind regards,

Catarina Salgado (on behalf of all authors)

References:

1. Lee DD, Leao R, Komosa M, Gallo M, Zhang CH, Lipman T, et al. DNA hypermethylation within TERT promoter upregulates TERT expression in cancer. The Journal of clinical investigation. 2019;129(4):1801.

2. Horn S, Figl A, Rachakonda PS, Fischer C, Sucker A, Gast A, et al. TERT promoter mutations in familial and sporadic melanoma. Science (New York, NY). 2013;339(6122):959-61.

3. Huang FW, Hodis E, Xu MJ, Kryukov GV, Chin L, Garraway LA. Highly recurrent TERT promoter mutations in human melanoma. Science (New York, NY). 2013;339(6122):957-9.

4. Schwartz U, Nemeth A, Diermeier S, Exler JH, Hansch S, Maldonado R, et al. Characterizing the nuclease accessibility of DNA in human cells to map higher order structures of chromatin. Nucleic acids research. 2019;47(3):1239-54.

5. Valouev A, Johnson SM, Boyd SD, Smith CL, Fire AZ, Sidow A. Determinants of nucleosome organization in primary human cells. Nature. 2011;474(7352):516-20.

6. Stern JL, Paucek RD, Huang FW, Ghandi M, Nwumeh R, Costello JC, et al. Allele-Specific DNA Methylation and Its Interplay with Repressive Histone Marks at Promoter-Mutant TERT Genes. Cell reports. 2017;21(13):3700-7.

7. Nell RJ, Steenderen Dv, Menger NV, Weitering TJ, Versluis M, van der Velden PA. Quantification of DNA methylation using methylation-sensitive restriction enzymes and multiplex digital PCR. 2019:816744.

8. Zhu J, Zhao Y, Wang S. Chromatin and epigenetic regulation of the telomerase reverse transcriptase gene. Protein & cell. 2010;1(1):22-32.

9. Xu D, Popov N, Hou M, Wang Q, Bjorkholm M, Gruber A, et al. Switch from Myc/Max to Mad1/Max binding and decrease in histone acetylation at the telomerase reverse transcriptase promoter during differentiation of HL60 cells. Proceedings of the National Academy of Sciences of the United States of America. 2001;98(7):3826-31.

10. Wang S, Hu C, Zhu J. Transcriptional silencing of a novel hTERT reporter locus during in vitro differentiation of mouse embryonic stem cells. Molecular biology of the cell. 2007;18(2):669-77.

11. Atkinson SP, Hoare SF, Glasspool RM, Keith WN. Lack of telomerase gene expression in alternative lengthening of telomere cells is associated with chromatin remodeling of the hTR and hTERT gene promoters. Cancer research. 2005;65(17):7585-90.

12. Liu C, Fang X, Ge Z, Jalink M, Kyo S, Bjorkholm M, et al. The telomerase reverse transcriptase (hTERT) gene is a direct target of the histone methyltransferase SMYD3. Cancer research. 2007;67(6):2626-31.

13. Zhu Q, Liu C, Ge Z, Fang X, Zhang X, Straat K, et al. Lysine-specific demethylase 1 (LSD1) Is required for the transcriptional repression of the telomerase reverse transcriptase (hTERT) gene. PloS one. 2008;3(1):e1446.

14. Devereux TR, Horikawa I, Anna CH, Annab LA, Afshari CA, Barrett JC. DNA methylation analysis of the promoter region of the human telomerase reverse transcriptase (hTERT) gene. Cancer research. 1999;59(24):6087-90.

15. Dessain SK, Yu H, Reddel RR, Beijersbergen RL, Weinberg RA. Methylation of the human telomerase gene CpG island. Cancer research. 2000;60(3):537-41.

16. Guilleret I, Yan P, Grange F, Braunschweig R, Bosman FT, Benhattar J. Hypermethylation of the human telomerase catalytic subunit (hTERT) gene correlates with telomerase activity. International journal of cancer. 2002;101(4):335-41.

17. Renaud S, Loukinov D, Bosman FT, Lobanenkov V, Benhattar J. CTCF binds the proximal exonic region of hTERT and inhibits its transcription. Nucleic acids research. 2005;33(21):6850-60.

18. Wang S, Zhu J. Evidence for a relief of repression mechanism for activation of the human telomerase reverse transcriptase promoter. The Journal of biological chemistry. 2003;278(21):18842-50.

19. Wang S, Zhu J. The hTERT gene is embedded in a nuclease-resistant chromatin domain. The Journal of biological chemistry. 2004;279(53):55401-10.

20. Woodcock CL, Ghosh RP. Chromatin higher-order structure and dynamics. Cold Spring Harbor perspectives in biology. 2010;2(5):a000596.

21. Ou HD, Phan S, Deerinck TJ, Thor A, Ellisman MH, O'Shea CC. ChromEMT: Visualizing 3D chromatin structure and compaction in interphase and mitotic cells. Science (New York, NY). 2017;357(6349).

22. Castelo-Branco P, Choufani S, Mack S, Gallagher D, Zhang C, Lipman T, et al. Methylation of the TERT promoter and risk stratification of childhood brain tumours: an integrative genomic and molecular study. The Lancet Oncology. 2013;14(6):534-42.

23. Barthel FP, Wei W, Tang M, Martinez-Ledesma E, Hu X, Amin SB, et al. Systematic analysis of telomere length and somatic alterations in 31 cancer types. Nature genetics. 2017;49(3):349-57.

Attachment

Submitted filename: Response to Reviewers.pdf

Click here for additional data file.^{(211.3KB, pdf)}

PLoS One. doi: 10.1371/journal.pone.0231418.r003

Decision Letter 1

Srinivas Saladi

24 Mar 2020

Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression

PONE-D-19-33667R1

Dear Dr. Salgado,

We are pleased to inform you that your manuscript has been judged scientifically suitable for publication and will be formally accepted for publication once it complies with all outstanding technical requirements.

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Reviewer #1: All comments have been addressed

Reviewer #2: All comments have been addressed

**********

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Reviewer #1: Yes

Reviewer #2: (No Response)

**********

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Reviewer #1: Yes

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**********

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**********

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**********

6. Review Comments to the Author

Reviewer #1: The authors have done a good job of addressing all the comments. They have explained the discrepancy between their results and previously published studies. They have also updated the methods section to be more comprehensive about the biochemical experiments and statistical tests performed.

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PLoS One. doi: 10.1371/journal.pone.0231418.r004

Acceptance letter

Srinivas Saladi

25 Mar 2020

PONE-D-19-33667R1

Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression

Dear Dr. Salgado:

I am pleased to inform you that your manuscript has been deemed suitable for publication in PLOS ONE. Congratulations! Your manuscript is now with our production department.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

S1 Table. Tailed primers used for amplification of 325-bp region in bisulfite-converted samples.

(XLSX)

Click here for additional data file.^{(9KB, xlsx)}

S2 Table. Primers and probe sequences to amplify the 106-bp amplicon in a novel design of a ddPCR assay to determine the methylation fraction.

(XLSX)

Click here for additional data file.^{(9KB, xlsx)}

S3 Table. Primers and probe sequences to amplify the 231-bp region encompassing 31 CpG sites around the cg11625005 in a novel ddPCR assay to assess the chromatin state.

(XLSX)

Click here for additional data file.^{(9.1KB, xlsx)}

S4 Table. Primer and probe sequences for TERT expression in qPCR.

(XLSX)

Click here for additional data file.^{(9.2KB, xlsx)}

S5 Table. Overview of the methylation fraction (measured by ddPCR and NGS), mutational status and TERT mRNA expression of our sample cohort (n = 61).

(XLSX)

Click here for additional data file.^{(17.5KB, xlsx)}

(XLSX)

Click here for additional data file.^{(14.4KB, xlsx)}

S7 Table. Raw data used in the Fig 5.

(XLSX)

Click here for additional data file.^{(9.2KB, xlsx)}

S8 Table. Raw data used in Fig 6.

(XLSX)

Click here for additional data file.^{(17.2KB, xlsx)}

S9 Table. Results overview.

(XLSX)

Click here for additional data file.^{(33.1KB, xlsx)}

Attachment

Submitted filename: Response to Reviewers.pdf

Click here for additional data file.^{(211.3KB, pdf)}

Data Availability Statement

All relevant data are within the manuscript and its Supporting Information file.

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PERMALINK

Interplay between TERT promoter mutations and methylation culminates in chromatin accessibility and TERT expression

Catarina Salgado

Celine Roelse

Rogier Nell

Nelleke Gruis

Remco van Doorn

Pieter van der Velden

Roles

Abstract

Introduction

Results

NGS-based deep bisulfite sequencing and development of a ddPCR assay to assess TERTp methylation fraction

Fig 1. Methylation fraction (MF) of 31 CpG sites around cg11625005 in 35 primary skin samples and 9 melanoma cell lines.

Fig 7. Results overview.

Fig 2. Methylation fraction (MF) analysed through ddPCR.

Absence of correlation between methylation fraction and TERT expression

Fig 3. Correlation between methylation fraction (%) and TERT mRNA expression in total of 31 samples: Fibroblasts (n = 3), melanocytes (n = 1), keratinocytes (n = 2), cutaneous melanoma cell lines (n = 19) and uveal melanoma cell lines (n = 6).

Evaluation of TERTp mutations in a collection of skin samples and melanoma cell lines

Fig 4. TERTp mutational status of primary skin samples and cutaneous melanoma cell lines.

Absence of correlation between mutational status and TERT expression

Fig 5. Correlation between TERTp mutational status and TERT mRNA expression in total of 31 samples: Fibroblasts (n = 3), melanocytes (n = 1), keratinocytes (n = 2), cutaneous melanoma cell lines (n = 19) and uveal melanoma cell lines (n = 6).

TERT expression is correlated to chromatin accessibility

Fig 6. Accessibility of TERTp around cg11625005 in 8 melanoma cell lines.

Discussion

Fig 8. Proposed model of TERT transcriptional regulation.

Material and methods

Samples, DNA extraction and PCR

Table 1. Samples overview.

Promoter methylation determination

Bisulfite conversion and next-generation sequencing (NGS)-based deep bisulfite sequencing

Novel design of a ddPCR assay using methylation-sensitive restriction enzymes (MSREs) to determine TERTp methylation fraction

Assessment of mutational status

Sanger sequencing

Mutation analysis using commercial TERT C250T and C228T mutation assays

Chromatin accessibility

Cell culture and treatment to assess chromatin states

Novel design of a ddPCR assay to assess chromatin opening state

Allele-specific chromatin accessibility

RNA isolation, cDNA synthesis and quantitative real-time PCR

Statistical analysis

Ethics statement

Supporting information

Acknowledgments

Data Availability

Funding Statement

References

Decision Letter 0

Srinivas Saladi

Roles

Transfer Alert

Author response to Decision Letter 0

Decision Letter 1

Srinivas Saladi

Roles

Acceptance letter

Srinivas Saladi

Roles

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

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