The adaptive molecular landscape of reprogrammed telomeric sequences

Domain: neuroscience
Published: 2026-04-09
Editorial outcome: major_revision

Abstract
Telomeric sequences vary across the tree of life and intimately co-evolve with telomere-binding protein complexes. However, the molecular mechanisms allowing organisms to adapt to new telomeric sequences are difficult to gauge from extant species. Here, we engineer multiple yeast lines to human-like telomeric repeats to unveil their molecular and fitness response to reprogrammed telomeres. Initially, the exchange of telomere sequences results in genome instability, proteome remodelling and severe fitness decline. Adaptive evolution experiments select for repeated mutations that drive adaptation to the humanized telomeres. These consist of the recurrent amplification of the telomere-binding protein gene TBF1, by complex aneuploidies, or in repeated mutations that attenuate the DNA damage response. Overall, our results outline a response that defines the adaptive molecular landscape to reprogrammed telomeric sequences.

Introduction
The correct maintenance of the genome is of fundamental importance in order to prevent instability and degeneration. In most eukaryotes, the genome is organized in linear segments whose extremities are exposed to degradation and need to be protected. This function is performed by ribonucleoproteins called “telomeres”, consisting of a repeated DNA sequence and its associated proteins1. Telomeres are maintained via the specialised enzyme telomerase. This enzyme is a reverse transcriptase that synthesizes new repeats using a RNA moiety as template2–4⁠.

Despite variations in their DNA sequence having occurred during evolution, the function of telomeres in genome maintenance is remarkably conserved across the tree of life. In fact, all telomeric DNA motifs share common characteristics, like being short, tandemly-repeated and G-rich, and the principles of telomere biology are widely conserved even among distant taxa5. Nevertheless, variations in both telomeric sequence and length occur6,7. The vertebrate-like TTAGGG (T AG ) repeat is the most widespread, being present in the majority of Metazoa, but also in some classes of protists, fungi and plants8. Variations of this motif are observed in other taxa, like TTTAGGG in plants9⁠, TTAGG in insects10, TTAGGC in nematodes11⁠ and degenerate motifs in Saccharomyces yeasts12. In other organisms, telomeric repeat units can be very long and complex, like in species of the genus Candida or in fission yeasts13,14. The vertebrate-like T AG repeat is thought to be the ancestral unit from which other variants evolved, given its wide-spread conservation across multiple phylogenetic lineages8. Current models of telomere sequence evolution involve the gradual modification of the telomerase RNA template followed by modifications of telomere-binding-proteins to accommodate the new telomeric sequences15,16. However, how organisms subsequently adapt to accommodate and potentially capitalize on these changes remains uncharacterised.

Telomeric DNA varies in length with a difference of several orders of magnitude across species and extensive within-species variation. Telomere length ranges from 20 bp in the ciliate Oxytricha trifallax17, 150-600 bp in budding yeasts18, to 5-15 kb in humans19 and over 100 kb in some mice species20. The budding yeast Saccharomyces cerevisiae has been a leading model in telomere biology. S. cerevisiae telomeres are constituted by degenerate TG repeats18⁠. Moreover, yeast telomeres are flanked by repetitive 1-3 subtelomeric DNA like the X and Y′ elements, and interstitial telomeric sequences (ITS) that are often located between them, adding further complexity18. S. cerevisiae cells can survive for several cell divisions after the complete loss of telomerase, and often generate stable survivors that can replicate indefinitely thanks to recombination-based maintenance of telomeres21.

It is possible to generate yeasts with human-like T AG telomeres by engineering the 2 3 telomerase RNA template encoded by the gene TLC122⁠. Such a humanization of yeast telomeres entails a switch in telomere-binding proteins, in which Rap1p, originally bound to native TG repeats, is replaced by Tbf1p, a regulatory factor known to bind TTAGGG 1-3 repeats in the yeast genome23,24. Humanized telomeres are dysfunctional and show deregulation of their length and loss of subtelomeric transcriptional silencing. Furthermore, they trigger the activation of the DNA damage response (DDR) with consequent cell cycle arrest25–33.

In this study, we investigate the evolutionary response of yeast cells to a sudden reprogramming of all telomeres by propagating them through experimental evolution. We sequentially combine two experimental evolution paradigms: first, we propagate cells over many mitotic generations through mutation accumulation lines (MAL) to minimize selection and study the long-term effects of telomere reprogramming. Next, we submit MAL to adaptive evolution (AEL) by repeated serial transfers of large populations in which multiple clones are competing against each other, to map mutations that counteract the fitness decline. The latter procedure ensures that if some clones develop a beneficial mutation, they will outcompete the others and lead the mutation to increase frequency.

During MAL, yeasts with humanized telomeres show a progressive fitness decay accompanied by genome instability and partial proteomic remodelling, but recover near-wild-type fitness after AEL by amplifying TBF1 or modulating the DNA damage response.

Results
An evolutionary framework of novel telomeres
Previous work has shown that S. cerevisiae yeasts engineered to carry point mutations in the TLC1-encoded 16 nt telomerase RNA template have growth and chromosome segregation defects34, suggesting that this motif is under strong evolutionary constraints. To quantify these constraints, we examined the template and its flanking regions in 8 Saccharomyces species spanning nearly 20 MY of evolution. Our multiple sequence alignment revealed that, while flanking regions show variation that reflects the species sequence divergence, the template region is extremely conserved, with only two variants: in the first and last nt positions (Fig. 1a). We further analyzed intraspecific variation in TLC1 in a collection of 3139 natural S. cerevisiae isolates to explore recent mutations across the ecological and geographic range of this model species. We found two rare variants, again in the first or the last nt of the template, with minor allele frequencies of 0.0003 (Fig. 1a). Thus, purifying selection acting on the TLC1 sequence in natural yeast lineages is remarkably strong.

In order to reconfigure S. cerevisiae telomere repeats, we used a genetic system that generates isogenic strains with either wild-type or humanized telomeres27,28⁠ (Supplementary Data 1). In brief, these two strains differ by a low-copy centromeric plasmid with either a native (pTLC1-y) or humanized (pTLC1-h) copy of TLC1, with the chromosomal TLC1 having been deleted from its native locus. These strains contain an additional plasmid with a copy of TLC1-y (pAZ1) that serves as a back-up and pAZ1 loss can be controlled by plating cells in a selective medium (Fig. 1b).

We initiated the telomere reprogramming by streaking the ancestors on 5-FOA for two single-cell bottlenecks (SCBs) to select for URA3-containing pAZ1 loss. We investigated the effect of pAZ1 loss on telomere DNA length and composition, as well as on mitotic cell growth and lifespan. First, we sequenced the ancestors’ genomes before (0 SCBs) and after pAZ1 loss (2 SCBs) to estimate telomere length and composition from short-read data35. The TG telomere length decreased upon pAZ1 loss, for both the wild-type 1-3 and humanized ancestor, consistent with telomere length being affected by the copy number of TLC136. The wild-type did not contain T AG repeats, while a small amount 2 3 was detected (5% T AG reads) in the humanized ancestor at both timepoints (Fig. 1c 2 3 and Supplementary Data 2-3). The rare T AG telomeric repeats and the loss of pAZ1 2 3 did not immediately affect the cell's doubling time across several conditions or chronological lifespan, nor the amount of repetitive subtelomeric elements (Fig. 1d-f, Supplementary Data 4-7).

Next, we investigated the impact of pAZ1 loss on the global proteome. We only detected 4 differentially expressed proteins in the humanized ancestor at 0 SCBs as compared to the wild-type (Supplementary Data 8-9). However, we identified 14 differentially expressed proteins in the humanized ancestor at 2 SCBs. In humanized yeasts, ADE2 is inactivated by a loss-of-function mutation in its native locus and a functional copy is inserted into the right subtelomere of chrV and acts as subtelomeric transcriptional silencing reporter. The replacement of Rap1p by Tbf1p results in transcriptional derepression of subtelomeric genes most likely due to the lack of the deacetylases Sir2-4 binding to the decreased number of telomeric Rap1p25 and/or the insulator activity of Tbf1p. Consistent with an early loss of transcriptional silencing, we found Ade2p to be upregulated in yeast cells carrying the humanized TLC1 at 2 SCBs (Fig. 1g, Supplementary Data 8-9). Overall, we found the controlled loss of pAZ1 to initiate the telomeric replacement of yeast TG with human T AG repeats and that this initiation 1-3 2 3 coincided with modest proteomic remodelling. Thus we conclude that this system can be used to study the long-term consequences of a telomeric repeat reprogramming.

Chromosome end structures of humanized yeasts
We induced the telomere reprogramming by evolving 16 clones from the wild-type and humanized ancestors for 100 SCBs through a mutation accumulation lines (MAL) protocol, which minimizes selective pressure (Fig. 2a, Supplementary Fig. 1 and Supplementary Data 1). During MAL, humanized yeasts’ TG repeats are gradually 1-3 replaced by T AG repeats, but they maintain a short centromere-proximal stretch of TG 2 3 1-3 repeats26. Five wild-type lines became “petites” (growth deficiency caused by mutation or loss of mitochondrial DNA) and were not further characterized. We estimated that the mutation accumulation lines underwent ~2140 and ~1640 generations for wild-type and humanized lines, respectively (Supplementary Data 1). Several wild-type and humanized lines subsequently were used to initiate adaptive evolution lines through 31 serial transfers (STs, ~200 generations) of large population sizes to explore adaptation to new telomeric sequences (Fig. 2a and Supplementary Fig. 1).

To characterize the telomere DNA upon reprogramming, we applied short-read and long-read technologies to both MAL and AEL. Short-read sequencing confirmed the absence of T AG repeats in wild-types, while their amount increased in the humanized lines over 2 3 mitotic generations (Fig. 2b and Supplementary Data 2). ITS content and Y’ copy number (CN) increased in humanized yeasts, a hallmark of type I survivors when cells overcome telomerase inactivation via recombination-mediated telomere lengthening. Humanized yeasts also carried ITS with T AG repeats suggesting telomere-subtelomere 2 3 recombinations (Fig. 2c-d and Supplementary Data 2).

Long-read sequencing can resolve telomeres at single chromosome-end resolution19,37. We thus applied Nanopore sequencing to the humanized ancestor at 2 SCBs, 4 MAL and 2 single clones from the same AEL (Fig. 2e). At 2 SCBs, three telomeres (chrVI_R, chrXI_L, chrXVI_L) contained small amounts of terminal T AG repeats, confirming that 2 3 telomeric reprogramming starts swiftly. At 100 SCBs, almost all telomeres are constituted by an internal TG (median 45.5±23.3 bp) and a terminal T AG stretch (median 1-3 2 3 224±357.5 bp). While the TG length is similar across telomeres, the T AG lengths 1-3 2 3 show remarkable variation with the Hum15 line with ultra-long telomeres (>3 kb in chrIV_R) (Fig. 2e, Supplementary Fig. 2a and Supplementary Data 3). The probability of a telomere to be elongated by telomerase increases as its length decreases. When a telomere approaches critical length, telomerase will elongate it to restore a normal length18,38. In the case of humanized yeasts, this implies that a minimum tract of TG 1-3 repeats will never get eroded and replaced by T AG repeats. Humanized yeasts' 2 3 telomeres are subject to a higher turnover27. This, together with the stochasticity of telomerase's action, explains a great part of the humanized yeasts' telomere heterogeneity.

ITS and Y' elements consistently increased in MAL and AEL, with chrVIII_L and chrXVI_R being preferential sites for their amplification (Fig. 2f). We confirmed the presence of T AG repeats at ITS, with 5 conformations, implying recombination between telomeres 2 3 and ITS (Fig. 2g, Supplementary Fig. 2b-c and Supplementary Data 3-4). In contrast, X elements were largely stable with 1 copy per chromosome-end, except for chrXVI_R that carried 2 copies and occasionally amplified on chrVIII_L (Fig. 2f and Supplementary Data 4). Abundant chromosome size variations that are compatible with subtelomeric amplifications were also observed (Supplementary Note 1 and Supplementary Fig. 2d). These results raise the possibility that the amplification of subtelomeric ITS and Y' elements compensates for the inadequate chromosome end protection in humanized yeasts. To further probe this hypothesis, we analyzed co-variation between telomere length, ITS and Y’ CN. Y’ and ITS CN were strongly correlated both in MAL and AEL (r=0.96 and r=0.99, p<2.2e-16) (Supplementary Fig. 2e).

The high rate of amplification of subtelomeric ITS and Y' elements is reminiscent of type I survivors, while the presence of ultra-long and fully humanized telomeres resembles type II survivors, which tend to have minor subtelomeric amplifications but extremely long telomeres18. The activation of these alternative-lengthening-of-telomeres mechanisms likely contributes to telomere length heterogeneity in humanized yeasts.

We investigated the impact of telomere humanization on proteins that directly bind telomeric repeats and the telomere silencing reporter. The abundance of Ade2p was significantly higher in humanized lines than in wild-types in both MAL and AEL, consistent with loss of transcriptional silencing in subtelomeres (p=0.0003 in MAL at 100 SCBs). This shift was mirrored by lower Rap1p levels (p=0.09 in AEL from 100 SCBs) and higher Tbf1p levels (p=0.02 in MAL at 100 SCBs), consistent with humanized yeasts adjusting the level of their telomere-binding-proteins according to telomere DNA sequence composition or length (Fig. 2h). In contrast, only 15 of the 147 known telomere-length-maintenance proteins that were present in our proteomics dataset showed significant difference between humanized and wild-type yeasts (Supplementary Note 2).

Overall, we disentangled the telomere and subtelomere structure of yeasts with humanized telomeres at unprecedented resolution, revealing high telomere/subtelomere dynamics and heterogeneity.

The fitness and molecular landscape of humanized yeasts
We investigated the impact of telomere humanization on yeast fitness by measuring mitotic cell growth and chronological lifespan. First, we measured growth in the same medium used in our experimental evolution by monitoring the optical density in liquid cultures over 72 hours. During MAL, the cell doubling time remained largely stable for the wild-types, while humanized lines showed early growth defects followed by partial recovery (median doubling time 2.0/2.8 h for WT/Hum at 100 SCBs, p=1.5e-7). This indicates that some adaptation occurred despite the MAL being bottle-necked to random single cells (or at least extremely small populations) (Fig. 3a, Supplementary Fig. 3a-b and Supplementary Data 5).

We next tested whether telomere humanization impacts cell growth across other environments that were not used in our experimental evolution, including different carbon sources and stresses, by using the automated microbial phenomics framework Scan-o-matic39, and observed that rapamycin and glycerol magnified the fitness defects of humanized yeasts (Fig. 3b and Supplementary Data 6). We further measured the survival rate across a 12-days incubation in a rich medium without refreshing. Humanized lines progressively decreased their survival rate without recovering it (median viability 51/21 % for WT/Hum at day 12 and 100 SCBs, p=1.7e-5) (Fig. 3c, Supplementary Fig. 3c and Supplementary Data 7).

Next, we characterized the mitotic growth of AEL started from MAL after 100 SCBs. During this adaptive evolution, we observed complete growth recovery (median doubling time 2.0/2.0 h for WT/Hum at 31 STs, p=0.85). This recovery was evident also in non-adaptation environments, including rapamycin and glycerol (p=0.05 combining all conditions). However, it is accompanied by a decrease in survival rate, consistent with the fact that lifespan is not under selection in this evolutionary regime (median viability 32/18 % for WT/Hum at day 12 and 31 STs, p=0.09) (Fig.s 3a-c, Supplementary Fig. 4a and Supplementary Data 5-7). When started from 2 SCBs, both wild-type and humanized AEL had the same growth and survival and even improved their growth in certain environments when compared to the starting clones. Thus, if telomere reprogramming and selection progress in concert, cells keep their fitness (Fig. 3a-c, Supplementary Fig. 4b and Supplementary Data 5-7).

We examined the proteomic profiles of humanized and wild-type yeasts and derived a list of 110 proteins whose abundance changed significantly in at least one evolution condition (MAL or AEL). We used hierarchical clustering to group proteins based on their log FC co-variation, uncovering 8 major clusters (Fig. 3e). Clusters 1,2,3,5 contain proteins down-regulated in humanized yeasts, with an enrichment for the pentose phosphate pathway/NADPH regeneration, transport/localization, aminoacid metabolism, and stress response, respectively. Proteins in clusters 1,5 remained downregulated after AEL from 100 SCBs, while those in clusters 2,3 returned to normal abundance. Of note, proteins in cluster 3 exhibit opposite trends between MAL and AEL from 2 SCBs. Cluster 4 contains proteins upregulated in humanized yeasts and is enriched for deoxyribonucleotides synthesis. Clusters 6 and 7 comprise proteins down- or upregulated in AEL from 2 SCBs, respectively. Cluster 8 contains proteins upregulated in all strains, with no functional enrichment (Fig. 3d and Supplementary Data 9). Proteomic profiles of humanized and wild-type yeasts were similar in AEL initiated from 2 SCBs, indicating that their differential protein abundance is mainly driven by growth advantage and faster metabolism, rather than telomere structure. This effect might underlie a common adaptation signature to the dropout medium through a modification of aminoacid metabolism. Furthermore, we performed Principal Component Analysis to group strains based on their proteomic profile and separated humanized and wild-type MAL. Humanized AEL from 100 SCBs clustered together with the wild type MAL and the ancestors, confirming that they recovered a proteomic profile similar to the wild types as they recovered their growth capacity. In contrast, both humanized and wild-type AEL from 2 SCBs clustered together (Fig. 3e).

We further defined a “Telomere Humanization Proteomic Response” (THPR) as the proteins that are differentially abundant in at least 2 humanized MAL at 40 or 100 SCBs (n=283). We compared the THPR with genes whose RNA is differentially expressed upon telomere dysfunction induced by a cdc13 temperature-sensitive mutation and that are present in our dataset (n=232), and found a significant overlap between them (n=79, two-tailed X2 test, p=6.5e-20, Supplementary Data 9)40. In contrast, we did not find a significant overlap between the THPR and Rap1p targets (n=345), indicating that the THPR is not driven by Rap1p-dependent transcription (Supplementary Note 2).

We further compared an early THPR (defined as the proteins that are differentially abundant in at least 2 humanized MAL at 40 SCBs, n=190), a late THPR (defined as the proteins that are differentially abundant in at least 2 humanized MAL at 100 SCBs, n=230) and a response specific to AEL (defined as the proteins that are differentially abundant in at least 2 humanized AEL from 100 SCBs, n=40). Through GO-term enrichment analysis, we found that upregulation of ribonucleotide reductase is an early response that persists even after AEL. Downregulation of trehalose and glycogen metabolism persists across MAL, while the early THPR is characterized specifically by downregulation of phospholipid metabolism and ceramide biosynthesis (Supplementary Fig. 5a, Supplementary Data 9 and Supplementary Note 2). We did not detect changes in expression of genes located near centromeres and the genes that were upregulated in the THPR do not seem to cluster in any genomic region.

Taken together, our experiments revealed that reconfiguring telomeric DNA sequences causes a severe fitness decline accompanied by a proteomic deregulation and partial reprogramming, but yeasts can recover wild-type growth.

Mutations in the DNA damage response rescue humanized yeasts fitness
We analyzed whole-genome sequencing data from MAL and AEL to characterize the mutational spectrum and genetic alterations underlying fitness recovery. We identified 250 new single-nucleotide variants (SNVs) in the 16 humanized MAL (49/51 % within subtelomeres/core), and 87 in the 11 wild-type MAL (17/83 % within subtelomeres/core). Given that subtelomeres cover 11% of the genome41 and assuming that selection had a negligible influence on variant distributions across MAL, we thus find that subtelomeres constitute a mutational hotspot in humanized yeasts (binomial test, p<0.0001). Humanized yeasts have significantly higher mutation rates than wild-types, which instead are in line with previous estimates42 (Fig. 4a, Supplementary Fig. 5b-c and Supplementary Data 10). A GO-term enrichment analysis revealed that wild-type MAL mutations were enriched in genes encoding ion binding proteins (p=0.03), while mutations in humanized MAL were enriched in genes encoding ceramide biosynthesis (p=0.10), cell periphery (p=0.06) and DNA damage response proteins (MAL Hum8 - TEL1 and MAL Hum4 - MEC1; p=0.09), suggesting that these pathways could underlie their fitness recovery (Fig. 4b and Supplementary Data 11). Mutations in ceramide biosynthesis genes have very low SIFT scores: mutations in LAC1 (SIFT=8e-5) and ISC1 (SIFT=0) are missense and located in conserved regions of the protein, and the mutation in LAG1 is a stop-gain, indicating that they are likely loss-of-functions (Supplementary Data 10). Ceramide acts as a signalling molecule and mediates cell cycle arrest, senescence and apoptosis43. Moreover, it can inhibit telomerase and induce telomere shortening in human cells44,45. Therefore, mutations that inhibit ceramide biosynthesis are likely adaptive. This is also in agreement with the downregulation of ceramide biosynthesis genes observed in humanized yeasts at 40 SCBs (Supplementary Data 9 and Supplementary Note 2).

Both the MEC1 (R1824) and TEL1 (L2308) mutations are aminoacid substitutions located in the FAT domain of the respective proteins, with the TEL1 mutation located in a highly conserved region and therefore predicted to be impactful. The humanized MAL carrying the TEL1 L2308R mutation underwent massive ITS/Y’ amplifications, chromosomal rearrangements and whole-genome duplication. The MEC1 R1824H mutation occurred between 40-60 SCBs whereas the TEL1 L2308R mutation occurred between 60-80 SCBs. The occurrence of TEL1 L2308R precedes fitness recovery, supporting that DDR inactivation rescues fitness in humanized yeasts (Fig. 4c, Supplementary Fig. 3a-b and Supplementary Data 5).

Bulk sequencing of AEL at 31 STs unveiled only few new mutations (n=13/38 in AEL starting from 2/100 SCBs), with the tryptophan tRNA genes recurrently affected (n=6). Mutations affecting components of the MRX complex, which binds DNA breaks or uncapped telomeres and then activates the DDR through the recruitment of Tel1p and Mec1p, occurred recurrently and independently in three AEL: Hum1 - MRE11 K477I; Hum2 - MRE11 Q447P; Hum3 - RAD50 L171P). These lines grew poorly at 0 STs but completely recovered their fitness after AEL (31 STs), and all three mutations are predicted impactful through SIFT score analysis (Fig. 4c). The two MRE11 mutations in the DNA binding domain and RAD50 mutation in the ATPase domain were all fixed, in line with that they are strongly beneficial (Fig. 4b-c and Supplementary Data 10-11). To confirm the adaptive role of the MRX mutations, we engineered MRE11 Q447P into the slow-growing humanized line 3 at 100 SCBs. This mutation produced a full growth recovery across conditions, but decreased the survival rate (Fig. 4d and Supplementary Data 5-7). We tested whether the beneficial effect would also manifest in fit humanized yeast that had not yet suffered a loss of reproductive capacity by introducing MRE11 Q447P in the humanized line 1 at 2 SCBs. However, we observed no improvement in growth or lifespan, consistent with that its benefits are strictly limited to compensating for the fitness loss arising from telomere reprogramming (Supplementary Fig. 5d and Supplementary Data 5-7). The full deletion of MRE11 improved both growth and survival rate but did not match the growth of the wild type MAL (Supplementary Fig. 5e and Supplementary Data 5-7). Taken together, these results suggest that MRE11 mutations induce partial loss-of-function.

To further disentangle how DDR modulation rescues fitness in humanized yeasts, we inspected protein abundance of known DDR targets in the proteomic dataset. We observed upregulation of the ribonucleotide reductase (RNR) subunits Rnr2p/Rnr4p and the deoxyuridine triphosphate diphosphatase Dut1p, and the downregulation of the RNR inhibitor Sml1p. The upregulation of Rnr2p/Rnr4p/Dut1p is highest in MAL at 40 SCBs while it decreases after AEL from 100 SCBs (p<10-2 for all proteins at 100 SCBs) (Fig. 4e). We further checked the abundance of Rnr2p/Rnr4p/Dut1p in a humanized AEL that spontaneously developed a mutation in MRE11 (Hum2 31 STs MRE11 Q447P), as well as a line where we introduced the same mutation by CRISPR/Cas9 genetic engineering (Hum3 100 SCBs MRE11 Q447P). The three proteins decrease when MRE11 Q447P is present, suggesting attenuation of the DDR (Fig. 4f).

Overall, these findings reveal that humanization of yeast telomeres increases mutation rate and the partial inactivation of DDR rescues their fitness through its downstream targets.

Chromosome XVI aneuploidy is adaptive in humanized yeasts
After revealing the beneficial effect of DDR point mutations, we investigated whether adaptation also occurred via large-scale copy number and structural variants (SVs). All the MAL remained haploid except for a whole-genome duplication in the humanized MAL 8 with a TEL1 loss-of-function mutation. This line likely underwent endoreduplication since the MAT locus remained homozygous (MATa/MATa) (Fig. 5a and Supplementary Data 12). We next analyzed long-read-sequencing-derived genome assemblies from 4 humanized MAL at 100 SCBs (Hum1,4,8,15) and an ancestor at 2 SCBs. We did not detect any major SVs except for Hum8 at 100 SCBs, which underwent a translocation between chromosomes V and XV mediated by the two ADE2 copies (Supplementary Fig. 6, Supplementary Data 12 and Supplementary Note 1).

We next analysed read coverage along chromosomes to determine whether large CNV and aneuploidies occurred during MAL . We discovered one segmental duplication of chromosome IV (~100 kb, WT2) and 17 whole chromosome aneuploidies in our lines. These were all chromosome gains and exclusively occurred in humanized lines (except one in WT6). Chromosome XVI was aneuploid in 12 lines suggesting a possible adaptive role (Fig. 5a and Supplementary Data 12). We further explored whether an additional chromosome XVI is beneficial by comparing humanized lines carrying 1 or 2 copies of this chromosome. Lines with 2 copies had shorter telomeres, and this difference was driven by fewer T AG repeats (p=0.01) (Fig. 5b, Supplementary Fig. 7a and 2 3 Supplementary Data 2). Furthermore, they had a shorter doubling time in multiple environments but lower survival rate (p=8.8e-8 for doubling time across conditions, p=0.02 for survival rate at day 12) (Fig. 5b, Supplementary Fig. 7b and Supplementary Data 5-7). To better pinpoint the emergence of the aneuploidy during MAL evolution, we probed its existence in the stored records through qPCR and found that it occurred between 60-100 SCBs (only one case at 40-60 SCBs), with its emergence correlating well with the decrease of both doubling time and survival rate, confirming a causative role (p=0.22/0.02 for doubling time/survival at day 12) (Fig. 5a, Supplementary Fig. 7c-e and Supplementary Data 5-7).

Aneuploidies are usually detrimental in nature, especially when involving long chromosomes, but can also fuel adaptation46–48. We inspected whether any telomere-binding-proteins were encoded on chromosome XVI, detecting Tbf1p, which directly binds T AG repeats23. We tested the effect of TBF1 duplication by engineering its copy 2 3 number, first by deleting one TBF1 copy in a humanized MAL carrying two chromosomes XVI (Hum9) and then by ectopically inserting a second TBF1 copy in the HO locus on chromosome IV in a humanized MAL carrying a single chromosome XVI (Hum12), to uncouple TBF1's dosage effect from chromosome XVI's dosage. We found that the addition of a second TBF1 copy to a humanized MAL carrying one chromosome XVI improved growth, while the deletion of one out of two TBF1 copies increased the doubling time by 17%, indicating that TBF1 alone can rescue growth in humanized yeasts (Fig. 5c). The beneficial effect of an extra TBF1 in humanized yeasts likely derives from better telomere capping and/or Tbf1p’s inhibitory action towards the binding of the MRX complex to telomeres and its subsequent activation of DDR, as supported in our proteomic data by the inverse relationship between Tbf1p levels and DDR targets (Rnr2p, Rnr4p, Dut1p)24,32 (Supplementary Fig. 8a-b). These results suggest that the fitness advantage conferred by TBF1 duplication is due to a combination of telomere protection and DDR-inhibitory functions.

Next, we investigated the evolutionary dynamics of large-scale structural variants during AEL. All the AEL derived from ancestors at 2 SCBs, i.e. from before substantial telomere reprogramming, maintained their original ploidy and showed no major structural variants. In contrast, AEL derived from MAL at 100 SCBs and that had gained extra chromosomes during the MAL evolution, frequently lost the extra chromosomes during the adaptive evolution and thereby restored a balanced genome content. Humanized lines 2,8,3 lost their additional copies of chromosomes V, XI and XVI, respectively, while Hum11 revealed a mixed population of clones carrying different segments of the extra copy of chromosome XVI (Fig. 5a and Supplementary Data 12). We isolated 8 single clones from this population and uncovered 4 distinct chromosome XVI conformations by qPCR and whole-genome sequencing. All the segments' breakpoints were flanked by repetitive sequences such as Ty and long terminal repeats that likely mediated the recombination events. Notably, the region encompassing the TBF1 gene remained duplicated in all conformations and included the centromere (CEN16) (Fig. 5d). We applied long-read sequencing to two of these clones to disentangle their chromosome XVI structure. Clone D carried almost the full version of duplicated chromosome XVI, except for a terminal segment, and the duplicated segment was stabilized by the duplication and translocation of chrVIII-R telomeres. Clone H carried only a central duplicated segment of chromosome XVI and this segment was circularized with the circularization likely mediated by Ty elements and stabilized by CEN16 (Fig. 5e). This circular structure is likely present in all conformations lacking terminal duplicated blocks (i.e. conformations 2,3,4) and has the advantage of not needing telomeres. We tested whether these segmental duplications resulted in a double amount of proteins. Proteomic profiles mirrored read coverage ones for the 4 chrXVI conformations (Supplementary Fig. 8c). However, the average abundance for proteins encoded by genes in these segments was less than 2-fold higher than that of other proteins, indicating a buffering effect. An extension of this analysis to the whole genome revealed a partial gene dosage compensation at the proteome level, which was probably selected during the long-term evolution (Fig. 5f and Supplementary Note 3)49. We reasoned that this conformations mixture might be the result of selection favouring conformations carrying the shortest duplicated block containing TBF1, in order to minimize the detrimental effects associated with the duplication of all chrXVI genes. To test this hypothesis we measured growth in the 8 clones and confirmed an inverse relationship between growth and chrXVI duplicated content (p=2.9e-5 between conformations 1-2 and 3-4) (Fig. 5g and Supplementary Data 5-7).

Taken together, these results show that chrXVI aneuploidy is a primary evolutionary route to fitness recovery driven by the duplication of TBF1.

Methods
Yeast strains and plasmids
The two ancestor strains (one for the humanized and one for the wild-type lines) derive from Bah et al., 2004 and their genetic background is: RWY12 (BY4741 Mata, ura3-52, lys2–801, ade2-101, trp1-Δ1, his3-Δ200, leu2-Δ1, tlc1Δ::LEU2, VR-ADE2-T)65. These strains are haploid and auxotrophic for uracil, lysine, tryptophan and histidine. The native ADE2 gene on chromosome XV is not functional and another copy has been inserted into the right subtelomere of chromosome V. ADE2 inactivation results in red colonies through the accumulation of purine precursors in the vacuole and can be used as a marker of transcriptional silencing at subtelomeric sites. The replacement of the TLC1 gene with an auxotrophic marker ensures that the strains cannot use their native RNA template to elongate telomeres.

In addition, the two ancestors contain two centromeric plasmids: pAZ1 and pTLC1. The pAZ1 plasmid derives from Dionne & Wellinger, 1996 and contains the wild-type version of TLC1 and the selection marker URA3. The pTLC1 plasmid backbone originally derives from pRS314 in Sikorski & Hieter, 198966 and contains the TLC1 gene and the selection marker TRP1. The TLC1 gene in the pTLC1 plasmid is different in the two ancestors: the wild-type ancestor carries the native pTLC1-y gene while the humanized ancestor carries a modified pTLC1-h gene in which the 16 template nucleotides (5'-CACCACACCCACACAC-3') have been replaced by a humanized counterpart (5'-CUAACCCU-3').

Mutation accumulation lines
The humanized and wild-type ancestors were revived from frozen stocks and plated on Leucine/Tryptophan/Uracil dropout medium to select for cells containing both the pAZ1 and pTLC1 plasmids. Plates were incubated for 3 days at 30 °C. After that, we streaked for single colonies on 5-FOA plates to retrieve cells that had lost the URA3-containing pAZ1 plasmid, and therefore rely only on the TLC1 copy present in the pTLC1-y/pTLC1-h plasmids. We selected 16 colonies derived from the wild-type ancestor and 16 derived from the humanized ancestor, which contained only the pTLC1-y and pTLC1-h plasmids, respectively. Since two streaking-for-single were performed during the pAZ1 exclusion procedure, we dated this timepoint as 2 single cell bottlenecks (SCBs). Previous studies showed that the replacement of TG with T AG repeats is gradual and slow and it only 1-3 2 3 reaches a steady state after 1200 generations27, thus we planned our MAL to last for ~2000 generations. Assuming that yeasts perform ~20 generations at every SCB, our planned MAL duration was 100 SCBs.

We propagated the 32 lines on Leucine/Tryptophan dropout solid medium in order to maintain cells with tlc1::LEU2 genotype and to retain pTLC1-y/pTLC1-h. Cells at the final timepoint (100 SCBs) and at four intermediate timepoints (20, 40, 60, 80 SCBs) were stored at -80 °C. Moreover, we stored 4 humanized and 2 wild-type lines at 2 SCBs, immediately after the loss of pAZ1.

To estimate the number of generations occurring every 72 h, three colonies derived from three independent humanized lines at every timepoint and one colony from a wild-type line at 2 and 100 SCBs were resuspended in 100 µl of sterile water and serially diluted. Twenty µl of each dilution were plated on solid YPD medium and incubated for 72 h at 30 °C. The number of colonies was manually counted in the plate with a suitable dilution and the number of generations (G) was estimated according to G=log (n×d), where n is 2 the number of cells on the plate and d is the corresponding dilution factor. The total number of generations occurred during the MAL protocol for the wild-type and humanized lines (T and T ) were calculated as: T =G×100 where G was averaged across the wt hum wt only representative line at 2 and 100 SCBs;
T =(G ×20)+(G ×20)+(G ×20)+(G ×20)+(G ×20) where G was averaged across hum 20 40 60 80 100 x the three representative lines at each timepoint. The humanized lines at 2 SCBs were not considered for the calculation.

Adaptive evolution
The 4 humanized and 2 wild-type lines at 2 SCBs, as well as 5 humanized lines and 2 wild-type lines derived from the MAL protocol (Hum1,2,3,8,11,WT2,7) were further submitted to adaptive evolution (AEL) by multiple serial transfers (STs) of large population sizes. Strains were inoculated in tubes containing 5 ml of Leucine/Tryptophan dropout liquid medium and incubated at 30 °C in a shaking incubator. Serial transfers were performed every 48h by diluting 50 μl of the culture into 5 ml of fresh Leucine/Tryptophan dropout medium (1:100 dilution). Since the fitness defect caused by humanized telomeres is very pronounced, we carried out a relatively long AEL (30 STs) to have a higher chance of recovering adapted lines. At 31 STs, we measured the growth rate of our evolved lines and their ancestors and we found that our humanized lines had improved their growth. Thus, we stopped AEL at that point. To estimate the number of generations occurring at every serial transfer, we used the formula G=log (DF), where DF=dilution factor (1:100). 2 This gave us an estimate of 6.64 generations per serial transfer, and 206 total generations for the AEL.

Estimation of the growth performance in Leucine/Tryptophan dropout
Strains were incubated two nights in 200 ul of liquid Leucine/Tryptophan dropout medium in a non-shaking incubator at 30 °C. The day after, strains were mixed and 2 ul of the mixed culture was resuspended in 200 ul of fresh Leucine/Tryptophan dropout liquid medium in a 96-well plate. The plate was incubated in a TECAN plate reader machine (Tecan, Infinite F200 Pro) at 30 °C for 72 hours and optical density (600 nm) was measured every 15 minutes during the experiment. Resulting data were analyzed using the software PRECOG, where the doubling time is calculated as the mean of the five highest slopes in the exponential growth phase67. We phenotyped 3 biological replicates for each strain. Given the high number of strains in this study, phenotyping was performed using several 96-well plates. To correct for the batch effect and compare data across plates, we included the humanized ancestor at 0 SCBs (MJD548) in all the plates and used it as internal control. We normalized the data in each plate by dividing by the average of the 3 control replicates in that plate. We then recalibrated all the measurements by multiplying for constant values: 1.97 for population doubling time, 5.82 for lag duration and 3.20 for yield. These values represent the average of the 3 control replicates in the first plate. Values reported in Supplementary Data 5 and in the figures are already recalibrated.

Estimation of the growth performance across multiple conditions
Growth across multiple conditions was measured using the high-throughput and automated microbial phenomics framework Scan-o-matic, which monitors colony sizes on agar plates using desktop scanners and derives growth curves from these images39. Strains were grown in 5 ml Synthetic Complete (SC) shaking liquid culture overnight. 100 ul from each of these cultures were resuspended into four 96-well plates, each with a different randomized layout. These four 96-well plates were then transferred onto a 1536 SC PLOS agar plate using the Rotor HDA Robot (Singer ltd). Each randomized layout was replicated 3 times onto the 1536 plate, resulting in a total of 12 biological replicates (24 for SC). Each plate also contained 384 positions with spatial controls. The spatial control used was a wild-type ancestor at 2 SCBs (MJD5). Due to the size differences of the colonies when transferring from liquid to solid, the 1536 solid plate was grown for 3 days and then replica-plated onto a new SC agar plate. This new plate is the preculture plate. After 3 days of growth on the preculture plate, the strains were transferred onto the plates with the different stressors and phenotyped. Lag duration, population doubling time and yield were extracted from the resulting growth curves using Scanomatic (version 2.2). Lag was calculated as the intercept between the tangent of the initial colony size (average of the first three timepoints) and the tangent from the timepoints used to calculate the growth rate. Population doubling time was calculated at the steepest incline of the growth curve, using five timepoints for a local regression. Yield was calculated as the difference between the initial colony size, as calculated for lag, and the final colony size (average of the last three timepoints). All measurements were calculated from the smoothened growth curves.

Estimation of survival rate
Strains were grown two nights in liquid Leucine/Tryptophan dropout medium, diluted 100x in 200 μl of fresh Leucine/Tryptophan dropout in a 96-well plate and incubated at 30 °C for 15 days. We measured viability at the beginning of the stationary phase when cultures reached saturation (3 days), and then every 4 days until day 15. Data are shown by days post-saturation, meaning that day 0 post-saturation corresponds to day 3 after the inoculation of the culture, while day 12 post-saturation corresponds to day 15 post-inoculation. At each time point, 5 μl of cells were transferred in 100 μl of staining solution (Phosphate-buffered saline-PBS + 3 μM propidium iodide + 200 nM YO-PRO-1) in a 96-well plate and incubated 10 minutes in the dark at 30 °C. Cell viability was measured by high-throughput flow cytometry on a Cytoflex machine, using the plate reader module. Cells were excited with the 561nm Yellow/Green laser. Fluorescence was read with the Y610-mCHERRY filter for propidium iodide and with the B610-ECDA filter for YO-PRO-1. Flow cytometry data were subsequently analysed using the CytExpert software⁠. A first gate was defined on an FSC-A versus SSC-A plot to distinguish the yeast population from debris. A second gate was applied on an FSC-H versus FSC-Width plot to discriminate singlets from yeast aggregates. Cell viability within the singlet population was subsequently assessed based on exclusion of Propidium iodide and YO-PRO-1. Propidium iodide and YO-PRO-1 are membrane-impermeable nucleic acid binding molecules that can only enter into dead cells whose cellular membrane is damaged. Non-fluorescent cells were counted as viable whereas fluorescent cells were counted as dead (Supplementary Fig. 8d). We measured one biological replicate with 3000 events per sample.

Ploidy estimation and pulsed-field gel electrophoresis
Ploidy was estimated for MAL at 2 and 100 SCBs and after AEL (at 31 STs) using a propidium iodide (PI) staining assay. Ploidy was estimated at all timepoints during the MAL protocol for humanized line 8, which showed a diploid genome content at 100 SCBs. Cells were incubated in 100 ul of Leucine/Tryptophan dropout liquid medium in a 96-well plate and left two nights at 30 °C. After that, 3 μl were incubated in 100 μl of cold 70% ethanol for 3 h at 4 °C, washed twice with PBS, and resuspended in 100 μl of staining solution (15 μM PI, 100 μg/ml RNase A, 0.1% v/v Triton-X, in PBS). After that, samples were incubated for 3 h at 37 °C in the dark. Samples were analyzed on a FACS-Calibur flow cytometer using the HTS module for processing 96-well plates. Cells were excited at 488 nM and fluorescence was collected with a FL2-A filter. The distributions of both FL2-A and FSC-H values were processed to find the two main density peaks, which correspond to the two cell populations in G1 and G2 phase of the cell cycle.

Whole chromosome size was estimated for MAL at 2 and 100 SCBs through Pulsed-Field Gel Electrophoresis. Strains were grown two nights in liquid Leucine/Tryptophan dropout medium. Samples were centrifuged and the pellet was resuspended in 1 ml EDTA 50 mM. Samples were centrifuged again and the pellet was resuspended in 100 ul EDTA 50 mM. To make plugs, we added 50 ul of SCE solution (1 ml SCE, 9 mg zymolase, 50 ul beta-mercaptoethanol) and 0.5 ml of LMP agarose 1% (in EDTA 0.125 M) to each sample, immediately pipetted it into plug formers and let it cool on ice. We transferred the plugs into eppendorf tubes, added 0.5 ml of EDTA solution (9 ml EDTA 0.5 M, 1 ml TRIS HCl 1 M, 0.5 ml beta-mercaptoethanol) and incubated at 37 °C for four hours. After that, we removed the EDTA solution, added 10 ml of proteinase solution (10 mg proteinase K, 9 ml EDTA 0.5 M, 1 ml sodium sarkosyl 10%, 0.1 ml RNAase at 0.1 mg/ml) and incubated at 37 °C overnight. The day after, we removed the proteinase solution, washed the plugs twice in TE 1X for one hour, and resuspended them in 0.5 ml of TRIS/EDTA solution (9 ml EDTA 0.5 M, 1 ml TRIS HCl 1 M). Samples were separated on a 1% agarose gel for 15 hours with a switch time of 60 seconds followed by 9 hours with a switch time of 90 seconds at 6 V/cm and 14 °C. The switch angle was 120.

Generation of MRE11 and TBF1 mutants
The deletion of MRE11 was performed by standard homologous recombination. The URA3-containing cassette was amplified from the laboratory S288C strain carrying a functional URA3 gene. 80-bp primers used for the amplification were designed to contain the first and last 20 bp of URA3 followed by 60 bp of homology with flanking regions upstream and downstream of MRE11. Samples were transformed using the standard lithium-acetate protocol, plated on selective medium lacking uracil and incubated at 30 °C for 3–7 days. Candidate transformed clones were validated by PCR using one primer designed inside URA3 and another primer designed on the outside region of MRE11.

The deletion of one copy of TBF1 in humanized strains carrying two chromosome XVI copies was performed by standard homologous recombination. The HYGMX-containing cassette was amplified from a plasmid carrying this antibiotic marker. 80-bp primers used for the amplification were designed to contain 20 bp of the MX region followed by 60 bp of homology with flanking regions upstream and downstream of TBF1. Samples were transformed using the standard lithium-acetate protocol, plated on selective medium containing hygromycin (200 ug/ml) and incubated at 30 °C for 3–7 days. Candidate transformed clones were validated by PCR using a combination of two PCR reactions: one reaction carried one primer designed inside HYGMX and another primer designed on the outside region of TBF1; the other reaction carried both primers designed inside TBF1. Only clones positive for both reactions were selected, as we wanted to delete only one copy of TBF1.

The introduction of MRE11 Q447P, as well as the ectopic expression of TBF1 in the HO locus on chromosome IV, were performed by using CRISPR/Cas9 genome editing. The plasmid harboring Cas9 was obtained from Addgene pUDP004 and linearized with BsaI. The resistance to acetamide was replaced with the resistance cassette to hygromycin which was amplified from a plasmid harboring the HYGMX gene. The gRNAs with the necessary nucleotides for self-cleavage were designed using the software UGENE and ordered as synthetic oligos from Eurofins Genomics (TM). The oligo was inserted in the plasmid using the Gibson assembly kit (NEB, Gibson Assembly®) with a ligation reaction of 1 hour at 50 °C. The plasmid was transformed into DH5-alpha competent bacteria and these were plated on ampicillin LB plates (100 µg/µl) and incubated overnight at 37 °C. Cells were screened by PCR to validate the correct assembly and positive colonies were inoculated in ampicillin LB broth (100 µg/uL) and incubated overnight at 37 °C. The plasmid was then extracted using the QIAprep Spin Miniprep Kit according to the manufacturer’s instructions. The cassette used for the engineering of MRE11 Q447P was designed to overlap the mutation site. The cutting site for Cas9 was modified to make it unrecognizable, by replacing one nucleotide with another one that did not change the aminoacid of the corresponding codon. The single-stranded parts of this cassette were ordered as a unique synthetic oligo at Eurofins Genomics. They were subsequently mixed at equimolar ratio, heated at 95 °C for 15 minutes and cooled down at room temperature to be used for the transformation. The cassette used for the knock-in of TBF1 into the HO locus was amplified from the humanized ancestor at 0 SCBs (MJD548). 80-bp primers used for the amplification were designed to contain the first and last 20 bp of TBF1 followed by 60 bp of homology with flanking regions upstream and downstream of HO.

Yeast samples were transformed using the standard lithium-acetate protocol. Cells were plated on selective medium containing hygromycin (200 μg/ml) and incubated at 30 °C for 3–7 days. MRE11 Q447P candidate transformed clones were validated by Sanger sequencing of selected colonies. ho::TBF1 candidate transformed clones were validated by PCR using one primer designed inside TBF1 and another primer designed on the outside region of HO. Positive clones were streaked on Leucine/Tryptophan dropout medium and grown for 2 days at 30 °C to allow plasmid loss and stored at -80 °C in 25% glycerol stocks. Primers used for these experiments are provided in Supplementary Data 13.

Quantitative PCR to track chrXVI copy number
To track the occurrence of chrXVI aneuploidy during MAL, we extracted DNA from yeast cells at various timepoints during the MAL protocol using a MasterPure Yeast DNA purification kit (Epicentre), following the manufacturer’s instructions. Primers were designed for the TBF1 locus and for one control locus (PWP2) on chromosome III, since this chromosome never underwent aneuploidy in any line. To determine the copy number of chrXVI blocks in 8 single clones derived from Hum11 at 31 STs (MJD460), we extracted DNA from the clones and designed one pair of primers for each block. Primers were designed on the following loci: FUM1-Block1; TBF1-Block2; RPC40-Block 3; NUT2-Block4. We also used the same control locus on chrIII (PWP2). Primers used for the qPCR are provided in Supplementary Data 13. DNA was diluted to 8 ng/ul for all the samples and the qPCR was performed using iTaq Universal SYBR Green Supermix (total volume: 10 µl) and run on an Applied Biosystems StepOne machine using the comparative ΔCt method. The PCR protocol was: initial denaturation (95 C; 15 min) followed by 45 cycles of: denaturation (95 C; 15 s), annealing (60 C; 30 s), extension (72 C; 30 s). We performed two technical replicates per sample per pair of primers. We calculated the copy number of the chrXVI regions covered by the primers by first subtracting the Ct of the chrIII control to all the primer pairs (ΔCt). We further normalized each primer pair's ΔCt by subtracting the corresponding ΔCt of an internal control strain that has one copy of each region (humanized ancestor at 0 SCBs-MJD548). We calculated the copy number (CN) of each region as CN=2-ΔΔCt. In addition to the investigated samples, we also included a humanized line aneuploid for the full length of chromosome XVI (Hum3-MJD420) and the humanized ancestor at 0 SCBs (MJD548) that has one copy of each chromosome, as positive and negative controls, respectively.

DNA extraction and short-read sequencing
Cells were inoculated in 5 ml of liquid Leucine/Tryptophan dropout medium and grown overnight at 30 °C in a shaking incubator. DNA was extracted using “Yeast Masterpure” kit (Epicentre, USA) following the manufacturer’s instructions. Illumina paired-end libraries (2×150 bp) were prepared according to manufacturer’s standard protocols and sequenced with an HiSeq 2500 instrument, at the NGS platform of Institut Curie. For the MAL experiment, two initial strains at 2 SCBs (one with the humanized background and one with the wild-type background) and all the strains at 100 SCBs were sequenced. For the AEL experiment, all the 5 humanized and 2 wild-type lines at 31 STs were sequenced. We additionally sequenced the humanized and wild-type ancestors at 0 SCBs, plus 8 single clones derived from Hum11 at 31 STs. DNA from these 10 samples was extracted using the same protocol as above, but sequencing was performed by Novogene (Cambridge, UK).

Long-read sequencing and structural variant analysis
Cells from a humanized line at 2 SCBs (Hum1), 4 humanized lines at 100 SCBs (Hum1,4,8,15) and two single clones derived from a AEL started from 100 SCBs (Hum11 at 31 STs) were grown for two nights in liquid Leucine/Tryptophan dropout medium at 30 °C. Genomic DNA was extracted using “Yeast Masterpure'' kit (Epicentre, USA) according to the manufacturer's instructions. No vortexing was performed during the DNA extraction to avoid excessive DNA fragmentation. The correct size distribution of DNA fragments after the DNA extraction was verified by gel electrophoresis using the lambda DNA as molecular weight size marker. The MINION (Oxford Nanopore) sequencing library was prepared using the SQK-LSK109 sequencing kit according to the manufacturer's instructions. The library was loaded onto a FLO-MIN106 flow cell and sequencing was run for 72 hours. Long read basecalling, demultiplexing and scaffolding were performed using the pipelines LRSDAY and LRSDAY_Patch (https://github.com/nicolo-tellini/LRSDAY-Patch) and the assembler was set as canu68. Dotplots were generated using nucmer and mummerplot. Structural variants were further confirmed by manually inspecting reads encompassing the breakpoints using IGV (v2.3.68).

Short-read data preprocessing and CNVs estimation
We mapped Illumina reads resulting from MAL and AEL on the S. cerevisiae SGD reference genome assembly using bwa mem (v0.7.12). Read mapping runs were stored in the Sequence Alignment/Map (SAM ) format and its binary form (BAM) and post-processing steps were performed by picard tools (v2.8.0). Sorting, indexing and per-position coverage calculation were performed using SAMtools (v1.2). Regions with higher coverage values as compared to the genome-wide median were considered as copy number variants and their effective copy number was calculated as the ratio between their median coverage and the median genome-wide coverage for the strain in which they were observed. Reads from each strain were mapped against the reference S288C genome from the SGD database using bwa mem, to obtain the baseline coverage. Coverages from the bam files were extracted using a custom script using the R library Rsamtools. We used the software freebayes (v0.9.5) to call markers from the BAM files. Results were stored in Variant Call Format (VCF) files. We filtered the VCF files to keep only SNVs/indels with quality higher than 20 and coverage higher than 10.

Identification of de novo SNVs in MAL and AEL
For MAL, we subtracted the SNVs/indels in the humanized strain at 2 SCBs (MJD1) from the humanized lines at 100 SCBs, and the SNVs/indels in the wild-type strain at 2 SCBs from the wild-type lines at 100 SCBs, using the vcf-isec program included in the suite VCFtools (v0.1.16). For AEL, we subtracted the SNVs/indels in the corresponding mutation accumulation lines at 100 SCBs from their derived AEL. Next, we searched for shared SNVs present in at least two MAL at 100 SCBs (or in two AEL for the AEL experiment) and removed them from the VCF files, based on the assumption that identical de novo SNVs are highly unlikely to arise independently and derive from incorrect mapping or poor coverage. All the remaining SNVs/indels were manually validated using IGV (v2.3.68). SNVs were annotated using the S. cerevisiae SGD genome assembly implemented in the online software Variant Effect Predictor (VEP). Further annotation of the functional impact of SNVs was carried out using mutfunc. We tested whether selection during MAL might have affected base substitution rates. The proportion of observed vs expected coding SNVs (74 vs 75 %) and non-synonymous SNVs (68 vs. 75 %) in wild-types supports a close-to-neutral mutation accumulation scenario (binomial test: p=ns), while the corresponding proportions in humanized yeasts (59 and 76 % for coding and non-synonymous mutations, respectively) indicate an effect of selection in this group, with mutations less likely to occur in coding regions (binomial test: p<0.0001). De novo variants derived from MAL were used to estimate base substitution and insertion/deletion rates (μ) as μ=(n/bp)/g, where n is the number of de novo SNVs present in each line, bp is the (haploid) genome size of the S. cerevisiae SGD genome assembly and g is the number of generations performed in the mutation accumulation experiment (2140 for wild-type lines and 1640 for humanized lines).

Experimental validation of de novo SNVs in MAL and AEL
Two SNVs from humanized MAL were validated by Sanger sequencing. One variant was located inside the gene MEC1 (Hum4: MEC1 R1824H) and the other was inside the gene TEL1 (Hum8: TEL1 L2308R). A pair of primers (upstream and downstream) was designed for each SNV using Unipro UGENE, and used to perform Polymerase Chain Reaction (PCR). We checked the presence of the mutations at all timepoints during the MAL protocol. PCR products were sequenced by Eurofins Genomics. The presence and the genotype of the variants were checked by visual inspection of the electropherograms. We confirmed the presence of both variants and discovered that the mutations in TEL1 occurred between 60 and 80 SCBs, while the mutation in MEC1 occurred between 40 and 60 SCBs. We used the same procedure to confirm the presence of MRE11 and RAD50 mutations in AEL started from 100 SCBs (Hum1: MRE11 K477I; Hum2: MRE11 Q447P; Hum3: RAD50 L171P). Primers used for these validations are provided in Supplementary Data 13.

Analysis of TLC1 variation
To analyze the extent of genetic variation in the TLC1 template and its flanking regions across the Saccharomyces sensu stricto species and across the S. cerevisiae population, we first downloaded published TLC1 sequences from Liti et al., 200669, spanning several S. cerevisiae and S. paradoxus isolates. We then downloaded the TLC1 sequence from the Saccharomyces Genome Database (SGD) and used it to search for orthologs in other Saccharomyces sensu stricto species in the NCBI database, using the blastn algorithm. We then performed a multiple sequence alignment using clustal omega (https://www.ebi.ac.uk/jdispatcher/msa/clustalo). Next, we gathered publicly available Illumina sequencing datasets relative to 3,139 S. cerevisiae strains. We run whole-genome short-reads alignments with bwa mem (v.0.7.17-r1198-dirty) and processed the bam files according to the latest samtools guidelines (samtools fixmate, sort, markdup and view) (v1.19). For each isolate, we produced a gVCF file using bcftools (mpileup, call, view and annotate) (v1.19) and extracted the genotypes for the 16 nt template (S288C v.R64-3-1_20210421, chrITI:308054-308069) and 20 nt upstream and downstream this sequence (chrII:308034-308053, chrII:308070-308089). We converted the gVCF genotypes in fasta sequences concatenating the alleles across consecutive positions with vcf2phylip.py (v.2.8) and created a multifasta file. The heterozygous positions were resolved according to the IUPAC nomenclature. At the end, we extracted the unique sequences, their occurrences and the list of isolates sharing the unique sequences with seqkit rmdup (v.2.4.0). The output was parsed with custom R scripts.

Telomere length, ITS content and Y' copy number estimation from short-read sequencing
Telomere length, ITS content and copy number of Y' elements were determined using YeaISTY35. Telomere length and ITS content were measured separately for the TG and 1-3 T AG motifs. For TG we used the published version of YeaISTY, while we developed 2 3 1-3 a new version compatible with the T AG motif for this study. We used a threshold of 20 2 3 bp of telomeric repeats stretch length for TG , and a threshold of 18 bp for T AG . By 1-3 2 3 comparing our results with the estimates derived from long-read genome assemblies, we determined that our lengths were underestimated and the underestimation was particularly pronounced for the T AG repeat. This is likely due to the fact that the T AG 2 3 2 3 motif is longer and more complex than TG and sequencing errors are more prone to 1-3 disrupt the continuity of T AG stretches. As a consequence, we avoid direct comparison 2 3 of total telomere lengths (TG +T AG ) between humanized and wild-type yeasts. 1-3 2 3

Subtelomeric elements annotations from long-read genome assemblies
X and Y' elements were annotated from long-read genome assemblies using LRSDAY68. Elements annotated as “partial” were further inspected by extracting the corresponding and surrounding genomic regions (~10 kb) with bedtools getfasta and by performing a blastn local alignment with the SGD reference genome. Most of the partial elements were instead full versions and were re-annotated using the coordinates derived from the local alignment, taking the longest possible matches as the start and end points of the elements.

Telomeres and ITS annotations from long-read genome assemblies
We annotated TG and TTAG repeats in the genome assemblies by using a custom Perl 1-3 2 3 script adapted from our recently published pipeline YeaISTY35. This script inspects the scaffolds of genome assemblies and detects single units of telomeric motifs (CA, CCA, CCCA, TG, TGG, TGGG). If units are consecutive to one another they are considered as part of the same telomeric repeats stretch and its start/end points are assigned, respectively, as the position of the first base of the first unit and the position of the last base of the last unit. Once we obtained a list of telomeric repeats stretches and their start/end points in the scaffolds, we merged the repeats which were interspaced by only 1 bp into a unique stretch by using the software mergeBed included in the suite bedtools (version 2.17.0) with the option “-d 1”. Then, we filtered the list to keep only the stretches longer than 10 bp and we manually inspected them to eliminate breaks that were likely due to sequencing errors. The length of the telomeric repeats stretch was then calculated as end-start + 1. The interstitial stretches were annotated as ITS if they closely preceded a Y' element, while the stretches at the termini were annotated as telomeres. Chromosome-ends for which we could not detect any telomeres due to insufficient sequencing coverage were classified as truncated and they appear as spots without bars in Figure 2e. We further mapped long reads derived from each strain onto its corresponding genome assembly using bwa mem (v0.7.12) and manually visualized long reads encompassing telomeres using IGV (v2.3.68).

Mass spectrometry and proteomics data analysis
Cultivation of library: strains were randomized manually in triplicates onto 96-well plates, with the exception of strains MJD436-447, MJD450-455, MJD481-500, and MJD586-597, each of which was cultivated once (187 samples). Strains were pinned onto solid Leucine/Tryptophan dropout agar using a Singer Rotor and incubated at 30 °C for 48 h. Subsequently, each colony was pinned into 200 µl of Leucine/Tryptophan liquid dropout medium and the samples were incubated at 30 °C without shaking. After 32 h, these pre-cultures were mixed thoroughly by pipetting and 80 µl of each sample were added to 1.55 ml of pre-warmed Leucine/Tryptophan liquid dropout medium in 96-well deep-well plates. One sterilized borosilicate glass bead was added per well in order to aid mixing. Plates were sealed with a Breathe-Easy membrane and incubated for 14.5 h at 800 rpm, 30 °C. Then, 1.4 ml of culture were transferred to a new deep-well plate, and cells were harvested by centrifugation (2880 xg, 5 min, 4 °C). The optical density at harvest was measured using an Infinite M Nano plate reader (Tecan). The supernatant after centrifugation was removed by inverting the plate, and cell pellets were immediately cooled on dry ice. Cell pellets were stored at -80 °C until lysis.

Proteomic sample preparation: samples for proteomics were prepared in a 96-well high-throughput format. Cells were lysed in 200 µl 100 mM ammonium bicarbonate, 7 M urea by beat beating using a Geno/Grinder (Spex). Subsequently, samples were reduced and alkylated using DTT (20 µl, 55 mM) and iodoacetamide (20 μl, 120 mM), respectively. Samples were then diluted by adding 460 µl of ml 100 mM ammonium bicarbonate. 500 µl per sample were digested using 2 µg Trypsin/LysC (Promega, Cat#V5072). After 17 h of incubation at 37 °C, 25 µl 20% formic acid were added to the samples, and peptides were purified using solid-phase extraction on C18 96-well plates. First, plates were prepared by conditioning with methanol (200 µl, 50 g, 1 min) and washing two times with 50% ACN (200 µl, 50 g, 1 min). Then, plates were equilibrated by three times washing with 3% ACN, 0.1% FA (200 µl, 50 g, 80 g, 100 g, respectively, 1 min each). The flow-through was discarded. Next, 200 µl of the digested samples were loaded (100 g, 1 min, with the flow-through discarded) and washed three times with 3% ACN, 0.1% FA (200 µl, 100 g, 1 min). After the last washing step, an additional centrifugation was performed to ensure all liquid had been pushed through the solid phase (180 g, 1 min). The cleaned peptides were eluted with 50% ACN in three steps (120 µl, 120 µl, 130 µl, 180 g, 1 min) into a 1.1 ml square well 96-well V-bottom collection plate. Eluted samples were vacuum-dried and subsequently reconstituted in 30 µl 0.1% formic acid. An equivoluminal pool of all samples (5 µl per sample) was generated to be used as technical controls during MS measurements. Peptide concentrations of samples and the pool were determined using a fluorimetric peptide assay kit (Thermo Scientific, Cat#23290). Samples with a too low optical density at harvest (OD600 < 0.25) and/or too low peptide concentrations (no peptide signal observed) were excluded from mass spectrometric measurements.

LC–MS/MS measurements: samples were analyzed in two batches (96 samples per batch plus analytical quality control (pool) injected every ten samples, 20 quality control samples in total) on a SCIEX ZenoTOF 7600 System mass spectrometer coupled to a Waters ACQUITY UPLC M-Class System. Prior to MS analysis, 200 ng peptides were chromatographically separated with a 60 min gradient on a Waters HSS T3 column (300um x 150mm, 1.8um) heated to 35 °C, using a flow rate of 5 µl/min where mobile phase A and B were 0.1% formic acid in water and 0.1% formic acid in acetonitrile, respectively. For data independent acquisition, a zenoSWATH MS/MS acquisition scheme with 85 variable size windows and 11 ms accumulation time was used. Ion source parameters were set to: ion source gas 1 & 2 – 12 and 60 psi respectively; curtain gas 25, CAD gas 7 and source temperature at 150°C; spray voltage – 4500V.

Data pre-processing: raw data were processed with DIA-NN (version 1.8, available from https://github.com/vdemichev/DiaNN) using the following settings: fragment ion m/z range set from 300-1800, mass accuracy set to 20 ppm at the MS2 and MS1 level, respectively, scan window set to 6, MBR (Match-between-runs) enabled, and Quantification strategy set as “Robust LC (high Precision)”. The spectral library was generated computationally based on a yeast reference proteome (UniProt UP000002311, accessed on 23.04.2021). The resulting report.tsv file was further pre-processed at the precursor level in R. First, all non-proteotypic peptides were excluded and precursors with q.values, PG.q.values, Global.q.values, Global.PG.q.values, or GG.q.values > 0.01 were excluded. Next, samples with very low precursor numbers (Z-score < -2.5) were removed. Batch correction was performed by equalizing the median precursor quantity of each batch. Protein quantities were determined for all proteins with two or more precursors mapped using the diann_maxlfq() function of the DIA-NN R package (https://github.com/vdemichev/diann-rpackage), resulting in a dataset covering 3503 proteins across 181 samples. No imputation was applied to the proteomic dataset and undetected proteins were treated as missing values.

Data analysis: after these pre-processing steps, we calculated the median of the three biological replicates and normalized them using the wild-type strain 1 at 2 SCBs (MJD5) as a reference, to quantify the relative changes. The quality of the biological replicates was confirmed by the high Pearson's correlation coefficients (>0.98) between each pair. To identify genes that are differentially expressed as compared to the reference, we calculated the fold-changes and the p-values corresponding to each gene using the edgeR library in R. The normalization factors for differences in measurement depth across different samples were calculated using the function calcNormFactors. Dispersion factors were estimated using the functions estimateCommonDisp, estimateTagwiseDisp and estimateTrendedDisp and we quantified the dispersion of the gene expression based on the mean values. The function exactTest was used to identify significant differential expressions, which were then visualized as volcano plots highlighting significant fold-changes in expression using the EnhancedVolcano function from the EnhancedVolcano library. The Principal Component Analysis was performed using the function fviz_pca_ind from the library factoextra in R.

Next, we derived lists of differentially abundant proteins for each strain by selecting those showing a log FC>=1 or log FC<=-1 and p<=0.05. Each list was further filtered on the 2 2 basis of the aneuploidies carried by each strain, in order to retain only proteins that are differentially abundant due to upregulated/downregulated expression and not by copy number modifications of their encoding genes located on duplicated chromosomes. Once we produced a list for each strain, we derived lists of differentially abundant proteins for humanized and wild-type yeasts by group (MAL at 40 and 100 SCBs, AEL from 2 and 100 SCBs) by selecting those present in >40% of the strains in each group and combined them, deriving 110 proteins whose abundance changed in at least one group as compared to the reference (Fig. 3d). In contrast, the list of proteins defining the THPR was derived by combining the proteins that are differentially abundant in at least 2 humanized MAL at 40 or 100 SCBs. The THPR was compared with a set of genes that are differentially expressed upon cdc13 temperature-sensitive mutation and were downloaded from Greenall et al., 2008, as well as with a list of Rap1p gene targets. The list of Rap1p targets was downloaded from SGD.

GO term analysis
Standard GO term analysis was performed separately on the variants extracted from wild-type and humanized MAL. Due to the small number of de novo variants occurred during AEL, we combined the variants in AEL from 2 and 100 SCBs and performed GO term analysis separately on the variants extracted from wild-type and humanized AEL. GO term analysis for de novo mutations was performed using the GO Term Finder tool available at Saccharomyces Genome Database (SGD). Significant GO terms were extracted by the algorithm implemented in the tool, with a False Discovery Rate (FDR) corrected α threshold of 0.1. GO term analysis for differentially abundant proteins was performed using the gprofiler2 package in Rstudio.

Statistics & Reproducibility
No statistical method was used to predetermine sample size, but we propagated the same number of lines for wild types and humanized yeasts for the MAL experiment (n=16 for each type). However, 5 wild type MALs became petite and were excluded from the analyses. The experiments were not randomized and the investigators were not blinded to allocation during experiments and outcome assessment. Statistical comparisons between humanized and wild type yeasts were performed using the Wilcoxon signed-rank test, unless stated otherwise.

Data availability
The genome sequencing data generated in this study have been deposited in the Sequence Read Archive (SRA) database, NCBI under accession codes BioProject ID PRJNA985049, Biosample ID SAMN35786841-SAMN35786899. The phenotyping and proteomics data are available at: https://github.com/mdangiolo89/Humanized_yeasts_project70 and within the supplementary information files. All the strains generated in this work are available upon request. Raw and processed mass spectrometry data have been deposited at the ProteomeXchange consortium via the PRIDE partner repository with the dataset identifier PXD064562 and are publicly available. The yeast reference proteome is available at UniProt under the accession number UP000002311. Source data are provided with this paper. The list of Rap1p targets is available at: https://github.com/mdangiolo89/Humanized_yeasts_project70.

Code availability
YeaISTY (Yeast ITS, Telomeres and Y′ elements estimator) and other custom scripts are available at: https://github.com/mdangiolo89/Humanized_yeasts_project70.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript, D’Angiolo and colleagues investigate the evolution of novel telomeric sequences by introducing human telomeric repeats (T₂AG₃) and the TLC1 gene encoding the telomerase RNA template into budding yeast. By combining mutation accumulation lines (MAL) and adaptive evolution lines (AEL), the authors elegantly follow how these “humanized” telomeres evolve over successive generations. The study convincingly shows that yeast progressively accumulate T₂AG₃ repeats, illustrating a clear transition from native to human telomeric architecture.\nImportantly, the authors demonstrate that the acquisition of humanized telomeres initially compromises cellular fitness and increases genome instability, leading to elevated mutation rates. Adaptive recovery is largely achieved through two routes: attenuation of DNA-damage checkpoint activity and the duplication of chromosome XVI, where increased gene dosage of TBF1 appears to be a key driver of fitness restoration.\nOverall, this work provides a compelling mechanistic and evolutionary perspective on how telomeric sequence composition shapes genome stability. The methodology is rigorous and diverse, the experimental effort substantial, and the conclusions well supported by the data. The manuscript is clearly written, well referenced, and of broad interest to the telomere, genome stability, and evolutionary biology fields. I support publication after the authors address the following points, which I believe will strengthen the manuscript and broaden its impact:\n1) The authors state in the introduction that telomeric sequence diversification occurred “multiple times during evolution.” Since plant and opisthokont telomeric sequences are highly conserved, this sentence could benefit from clarification and, ideally, a reference specifying whether these changes reflect independent evolutionary events across clades, or turnover within particular lineages such as S. cerevisiae. Adding specificity would strengthen the argument.\n2) It would be helpful to highlight additional key genes (e.g., ADE2) directly in the volcano plot (Figure 1) to facilitate interpretation without needing to refer to supplementary tables. Additionally, in some supplemental figures (e.g., Supplementary Fig. 1), the font appears quite small for print; if possible, increasing font size would improve readability.\n3) The enrichment of genes associated with ceramide biosynthesis and the cell periphery in humanized MAL strains is mentioned but not discussed. While the focus on DNA-damage response attenuation is appropriate and well explored, a brief comment on how alterations in membrane-associated pathways might relate to telomere stress or adaptive responses would be valuable, even if speculative.\n4) Please clarify (if possible) whether increased TBF1 dosage on chromosomes other than chromosome XVI (e.g., via local duplication or ectopic integration) would be sufficient for adaptation, or whether ChrXVI aneuploidy specifically provides an additional selective advantage.\n5) Given the genome-wide instability observed, do the authors detect changes in pericentromeric/centromeric silencing or increased frequency of neocentromeres in humanized yeast after multiple generations? Even if negative, commenting on this would be informative for the broader chromosome biology community.", "reviewer_2": "The manuscript D'Angiolo et al. presents a thorough and insightful investigation into how yeast cells respond—both adaptively and non-adaptively—to the replacement of their native telomeric sequences with human-like repeats. The study builds elegantly on previous observations, reinforcing and extending our understanding that telomere humanization triggers a DNA damage response (DDR), and that attenuation of this response—through specific mutations—leads to improved cellular fitness. Importantly, the authors find an increased genome-wide mutation rate in humanized cells. Furthermore, they identify aneuploidy involving amplification of the telomere-binding protein TBF1 as an early and recurrent adaptive mechanism. With a clever experimental design combining state-of-the-art proteomics, fitness assays, and long-term evolution experiments, the authors illuminate the molecular routes available to cells facing a sudden change in telomeric repeats and the telomeric instability associated with it. Overall, this study makes a substantial contribution to our understanding of how telomeric sequences and their associated protein networks can evolve under selective pressure, offering valuable insights into the evolutionary plasticity of chromosome end maintenance.\nMajor Comments\n• I’m unclear about the interpretation of the proteomic data presented in Figure 1g. The authors suggest these changes result from derepression of genes following Rap1 replacement with Tbf1 (line 134). However, only ADE2 appears significantly upregulated, while the other 13 genes show reduced expression compared to WT after 2 SCBs. Could the authors clarify this discrepancy?\n• The authors define a “telomere humanization proteomic response” (THPR) based on proteins differentially abundant in at least two humanized MALs (line 285). I assume the intent was to capture a response to extended human telomeric repeats, distinguishing it from an acute or adaptive response. However, MALs are also undergoing adaptation (as shown by the doubling time changes in Fig. 3a), raising questions about the purity of this response. In this context, using samples at 40 SCBs might better represent early telomere-specific responses than those at 100 SCBs. Additionally, many AEL lines retain substantial human telomeric repeats after evolution. Have the authors assessed whether the THPR is still present in these lines, and to what extent? Can the authors distinguish between a physiological response (most likely more prominent in the 40SCBs, and an adaptive one? More evident in the AELs, with some potential overlap with the 100SCB samples?\n• The discussion presents DDR attenuation and aneuploidy (via TBF1 amplification) as alternative adaptive strategies to humanized telomeres, potentially operating through the same mechanism (i.e., attenuating DDR and promoting faster division). The authors propose a two-step model—aneuploidy arising first, then being replaced by less costly DDR mutations—consistent with existing literature.\nHowever, evidence for this is dispersed across the manuscript, making it difficult to track which evolved lines follow which adaptation path. A visual summary showing which lines exhibit TBF1 amplification and which have DDR mutations would clarify the narrative. If no longitudinal data per line is available, the authors should explicitly state that the model of substitution is speculative.\nMinor Comments\n• The four differentially expressed proteins in the humanized ancestor at 0 SCBs are referred to in Table S11, but should be Table S12.\n• In line 166, the authors approximate the length of the AEL to ~400 generations. However, in the methods, they report 31 1:100 dilutions. This should correspond to 6.6 generations per transfer, resulting in ~200 generations. Is this a typo, or does the difference derive from a different estimate of the generations, and if so, which one?\n• In the text and figures, serial transfers are referred to as ‘st’ (line 166); however, to avoid confusion, I believe they should be referred to with capital letters, as for the SCBs (ST for singular, STs for plural, as present in the methods).", "reviewer_3": "This study integrates mutation accumulation (MAL) with adaptive evolution (AEL) to chart the adaptive molecular landscape triggered by telomeric sequence reprogramming. The authors document early genome instability, proteome-wide reprogramming, and fitness decline, followed by recovery via two recurrent routes: (i) increased TBF1 dosage—often through chrXVI aneuploidy and finer structural variants—and (ii) attenuating mutations in the DNA damage response (DDR). The work addresses a fundamental question—how telomere sequence changes drive long-term organismal adaptation—with a coherent multi-omics framework and notable novelty. The findings are of high conceptual interest and merit publication after additional validation.\nMajor concerns：\n1. The data presented in Figure 2E, which shows the proportion of T2AG3 sequences across chromosomes, is particularly interesting. We noted some variability in the humanization efficiency among different clones and chromosomes, an observation that could be further highlighted and discussed in the main text. To enhance the impact of this work, it would be helpful if the authors could elaborate on the potential biological or technical reasons behind the observed heterogeneity. Additionally, some discussion on the challenges of achieving complete humanization across an entire chromosome would be very insightful for the reader community.\n2. There are certain differences between human and yeast telomere-associated proteins. Would overexpressing some human telomere proteins in yeast make the study of humanized telomeres in yeast more meaningful?\n3. The authors employed two laboratory evolution protocols—mutation accumulation lines (MAL) and adaptive evolution lines (AEL)—to effectively investigate the evolution of telomere sequences. It is suggested that the authors clarify in the Discussion section the distinct mutational outcomes associated with each evolution method and provide a rationale for these differences. For instance, while the authors observed chromosome XVI aneuploidy specifically in the MAL group, other studies have indicated that aneuploidy adaptations are more commonly associated with adaptive evolution lines.\nMinor concerns:\n4. The key conclusions rely on assembly and quantification of terminal repeats using long reads. It is recommended that the Methods section be written in more detail.\n5. The key conclusions rely on changes in doubling time; the Results section should add the calculation methods, and explain why correlations were computed on the log scale or not log scale.\n6. The combination of mutation accumulation (MAL) and adaptive evolution (AEL) protocols is a powerful design that effectively distinguishes stochastic mutations from adaptive ones. However, the rationale for selecting the specific endpoints of 100 single-cell bottlenecks for MAL and 31 serial transfers for AEL is not explicitly justified. It would be important to clarify whether these thresholds were defined based on preliminary biological observations or statistical considerations.\n7. Line 257 mentions that although yeast mitosis has been restored during evolution, chronological lifespan has not been restored. Mechanism behind this could be further discussed.\n8. The discussion could be strengthened by connecting telomere sequence evolution with the broader concept of cross-species telomere complex compatibility. Since telomere maintenance relies on co-evolution between telomeric DNA and binding proteins, contrasting the yeast humanization model with examples from other taxa would help contextualize the evolutionary adaptability and constraints observed here.\n9. Petite mutants in wild-type MALs are discarded without quantifying their incidence in humanized lines or exploring telomere–mitochondria interactions."}, "author_response": "Reviewer #1 (Remarks to the Author):\nIn this manuscript, D’Angiolo and colleagues investigate the evolution of novel telomeric\nsequences by introducing human telomeric repeats (T₂AG₃) and the TLC1 gene encoding the\ntelomerase RNA template into budding yeast. By combining mutation accumulation lines (MAL)\nand adaptive evolution lines (AEL), the authors elegantly follow how these “humanized” telomeres\nevolve over successive generations. The study convincingly shows that yeast progressively\naccumulate T₂AG₃ repeats, illustrating a clear transition from native to human telomeric\narchitecture.\nImportantly, the authors demonstrate that the acquisition of humanized telomeres initially\ncompromises cellular fitness and increases genome instability, leading to elevated mutation rates.\nAdaptive recovery is largely achieved through two routes: attenuation of DNA-damage checkpoint\nactivity and the duplication of chromosome XVI, where increased gene dosage of TBF1 appears to\nbe a key driver of fitness restoration.\nOverall, this work provides a compelling mechanistic and evolutionary perspective on how\ntelomeric sequence composition shapes genome stability. The methodology is rigorous and diverse,\nthe experimental effort substantial, and the conclusions well supported by the data. The manuscript\nis clearly written, well referenced, and of broad interest to the telomere, genome stability, and\nevolutionary biology fields. I support publication after the authors address the following points,\nwhich I believe will strengthen the manuscript and broaden its impact:\n1) The authors state in the introduction that telomeric sequence diversification occurred “multiple\ntimes during evolution.” Since plant and opisthokont telomeric sequences are highly conserved, this\nsentence could benefit from clarification and, ideally, a reference specifying whether these changes\nreflect independent evolutionary events across clades, or turnover within particular lineages such as\nS. cerevisiae. Adding specificity would strengthen the argument.\n[R] We apologize if this was unclear. Extensive investigations to detect telomeric sequences of\nvarious taxa in published genomic sequences have led to the hypothesis that the vertebrate-like\nT AG repeat is the ancestral unit from which other variants evolved, given its wide-spread\n2 3\nconservation across multiple phylogenetic lineages, including basal branches (Fulnečková et al.,\n2013). Starting from this assumption, other studies have hypothesised models in which variations in\nthe telomeric sequences originate from the gradual accumulation of mutations in the gene coding\nfor the telomerase RNA template. Subsequently, the new telomeric repeats are accommodated by\nmodifications of telomere-binding-proteins (Červenák et al., 2021).\n[A] We rephrased this sentence in the introduction and clarified the current model of telomere\nsequence evolution by expanding this section (page 3, lines 50-57).\n2) It would be helpful to highlight additional key genes (e.g., ADE2) directly in the volcano plot\n(Figure 1) to facilitate interpretation without needing to refer to supplementary tables. Additionally,\nin some supplemental figures (e.g., Supplementary Fig. 1), the font appears quite small for print; if\npossible, increasing font size would improve readability.\n[R&A] We indicated the other 13 genes that are differentially expressed in the humanized ancestor\nat 2 SCBs in Figure 1g. Furthermore, we increased the font size in the figures to meet the figure\npreparation guidelines.\n3) The enrichment of genes associated with ceramide biosynthesis and the cell periphery in\nhumanized MAL strains is mentioned but not discussed. While the focus on DNA-damage response\nattenuation is appropriate and well explored, a brief comment on how alterations in membrane-\nassociated pathways might relate to telomere stress or adaptive responses would be valuable, even if\nspeculative.\n[R] Mutations in ceramide biosynthesis genes (LAC1, LAG1, ISC1) occurred and were maintained\nin three independent humanized MAL. LAC1 and LAG1 are components of ceramide synthase, an\nenzyme that synthesizes ceramide from C26(acyl)-coenzyme A and dihydrosphingosine or\nphytosphingosine, while ISC1 codes for inositol phosphosphingolipid phospholipase C, an enzyme\nthat hydrolyzes complex sphingolipids to produce ceramide. All three mutations have low SIFT\nscores: mutations in LAC1 (SIFT=8e-5) and ISC1 (SIFT=0) are located in conserved regions of the\nprotein, and the mutation in LAG1 is a stop-gain, indicating that they are likely to induce loss-of-\nfunction in the resulting proteins (Tables S10-11). Ceramide acts as a signalling molecule and\nmediates cell cycle arrest, cell senescence and apoptosis, both in yeast and in mammals (Hannun,\n1996; Hannun & Luberto, 2000). Moreover, ceramide can inhibit telomerase and induce telomere\nshortening in human cells (Ogretmen et al., 2001; Sundararaj et al., 2004). Therefore, mutations that\ninhibit ceramide biosynthesis are likely to have a beneficial effect on humanized yeasts and should\nbe under positive selection. This is also in agreement with our proteomics results in which we\nidentified downregulation of ceramide biosynthesis pathways in humanized yeasts at 40 SCBs (see\nresponse to point 7 raised by reviewer 2).\n[A] We now discuss this aspect in the results section (page 17, lines 372-382).\n4) Please clarify (if possible) whether increased TBF1 dosage on chromosomes other than\nchromosome XVI (e.g., via local duplication or ectopic integration) would be sufficient for\nadaptation, or whether ChrXVI aneuploidy specifically provides an additional selective advantage.\n[R] We apologize if this was unclear. We ectopically integrated an additional copy of TBF1 in the\nHO locus on chromosome IV in a humanized strain carrying only one chromosome XVI (and one\ncopy of TBF1), as illustrated in Figure 5c and explained in the methods (page 36, lines 906-930).\nThis was sufficient to decrease the doubling time of the strain carrying this construct by about 30\nminutes. Since this fitness gain was independent from a gain of copy of chromosome XVI, an\nincreased TBF1 dosage alone can rescue the growth in humanized yeasts. However, we cannot\nexclude that other genetic or epigenetic factors on chromosome XVI might also contribute to the\nfitness gain.\n[A] We clarified in the main text (pages 20-21, lines 477-485) that our TBF1 copy number increase\nresults from an ectopic integration of the gene in the HO locus on chromosome IV, without\nchromosome XVI duplication.\n5) Given the genome-wide instability observed, do the authors detect changes in\npericentromeric/centromeric silencing or increased frequency of neocentromeres in humanized\nyeast after multiple generations? Even if negative, commenting on this would be informative for the\nbroader chromosome biology community.\n[R] We did not detect changes in expression of genes located near centromeres and the genes that\nare differentially expressed in humanized yeasts do not seem to cluster in any genomic region.\nMoreover, S. cerevisiae has point centromeres and we do not expect to observe centromere\ninstability. We re-examined the genome assemblies of humanized yeasts derived from long-read\nsequencing and did not detect any cases of neo-centromere formation. On the other hand, all forms\nof extra chromosome XVI included a centromere that is required to ensure segregation. The\nassembly of Hum8 merits a separate discussion as it revealed a chromosomal rearrangement\ninvolving chromosomes V and XV. This rearrangement resulted in the internalization of chrV-R\ntelomeres as ITS and it is stabilized by chrXV-L telomeres and its centromere (CEN15). It is\nunclear whether the remaining parts of chromosomes V and XV are attached to each other or\nconstitute two independent chromosomes. The first scenario is more likely as the resulting\nrecombinant chromosome would be stabilized by chromosome V centromere (CEN5). The second\nscenario would involve the formation of a neo-centromere and is more unlikely, in fact we did not\ndetect any extra centromeres.\n[A] We added these results on page 14, lines 327-329 and in supplementary discussion 1 (page 2,\nlines 33-34).\nReviewer #2 (Remarks to the Author):\nThe manuscript D'Angiolo et al. presents a thorough and insightful investigation into how yeast\ncells respond—both adaptively and non-adaptively—to the replacement of their native telomeric\nsequences with human-like repeats. The study builds elegantly on previous observations,\nreinforcing and extending our understanding that telomere humanization triggers a DNA damage\nresponse (DDR), and that attenuation of this response—through specific mutations—leads to\nimproved cellular fitness. Importantly, the authors find an increased genome-wide mutation rate in\nhumanized cells. Furthermore, they identify aneuploidy involving amplification of the telomere-\nbinding protein TBF1 as an early and recurrent adaptive mechanism. With a clever experimental\ndesign combining state-of-the-art proteomics, fitness assays, and long-term evolution experiments,\nthe authors illuminate the molecular routes available to cells facing a sudden change in telomeric\nrepeats and the telomeric instability associated with it. Overall, this study makes a substantial\ncontribution to our understanding of how telomeric sequences and their associated protein networks\ncan evolve under selective pressure, offering valuable insights into the evolutionary plasticity of\nchromosome end maintenance.\nMajor Comments\n6) I’m unclear about the interpretation of the proteomic data presented in Figure 1g. The authors\nsuggest these changes result from derepression of genes following Rap1 replacement with Tbf1\n(line 134). However, only ADE2 appears significantly upregulated, while the other 13 genes show\nreduced expression compared to WT after 2 SCBs. Could the authors clarify this discrepancy?\n[R&A] We agree with the reviewer that it is unlikely that Rap1p's extrusion from humanized\ntelomeres is the reason of the downregulation of 13 genes in the humanized ancestor at 2 SCBs, as\nRap1p acts mainly as a transcriptional activator. We removed that sentence.\n7) The authors define a “telomere humanization proteomic response” (THPR) based on proteins\ndifferentially abundant in at least two humanized MALs (line 285). I assume the intent was to\ncapture a response to extended human telomeric repeats, distinguishing it from an acute or adaptive\nresponse. However, MALs are also undergoing adaptation (as shown by the doubling time changes\nin Fig. 3a), raising questions about the purity of this response. In this context, using samples at 40\nSCBs might better represent early telomere-specific responses than those at 100 SCBs. Additionally,\nmany AEL lines retain substantial human telomeric repeats after evolution. Have the authors\nassessed whether the THPR is still present in these lines, and to what extent? Can the authors\ndistinguish between a physiological response (most likely more prominent in the 40 SCBs, and an\nadaptive one? More evident in the AELs, with some potential overlap with the 100 SCB samples?\n[R] We thank the reviewer for this suggestion. We further defined an early THPR (proteins that are\ndifferentially abundant in at least 2 humanized MAL at 40 SCBs), late THPR (proteins that are\ndifferentially abundant in at least 2 humanized MAL at 100 SCBs) and AEL response (proteins that\nare differentially abundant in at least 2 humanized AEL from 100 SCBs). By intersecting these\nresponses and conducting a GO-term enrichment analysis, we found that the upregulation of\nribonucleotide reductase is present in both early and late THPR and persists even after AEL.\nDownregulation of trehalose and glycogen metabolism persists across early and late THPR but is\nlost upon AEL. Interestingly, the early THPR is characterized specifically by downregulation of\nphospholipid metabolism and ceramide biosynthesis, but this enrichment is lost in the late THPR.\nLoss-of-function mutations in ceramide biosynthesis genes (LAC1, LAG1, ISC1) occurred and were\nmaintained in three independent humanized MAL (Tables S10-11, see response to point 3 raised by\nreviewer 1). Ceramide acts as a signalling molecule and mediates cell cycle arrest, cell senescence\nand apoptosis, both in yeast and in mammals (Hannun, 1996; Hannun & Luberto, 2000). Moreover,\nceramide can inhibit telomerase and induce telomere shortening in human cells (Ogretmen et al.,\n2001; Sundararaj et al., 2004). Taken together, these results suggest that the early telomere-specific\nresponse already has an adaptive component, and downregulation or loss-of-function mutations in\nceramide biosynthesis genes can rescue the growth defects induced by telomere humanization.\n[A] We added a summary of these results in the main text (page 13, lines 318-326) and we\nextensively present them in supplementary discussion 2 (pages 2-3, lines 47-67). The early and late\nTHPR, AEL response and their intersections, as well as their GO-term enrichments, are shown in\nTable S9 and Supplementary Figure 5a.\n8) The discussion presents DDR attenuation and aneuploidy (via TBF1 amplification) as alternative\nadaptive strategies to humanized telomeres, potentially operating through the same mechanism (i.e.,\nattenuating DDR and promoting faster division). The authors propose a two-step model—\naneuploidy arising first, then being replaced by less costly DDR mutations—consistent with\nexisting literature. However, evidence for this is dispersed across the manuscript, making it difficult\nto track which evolved lines follow which adaptation path. A visual summary showing which lines\nexhibit TBF1 amplification and which have DDR mutations would clarify the narrative. If no\nlongitudinal data per line is available, the authors should explicitly state that the model of\nsubstitution is speculative.\n[R&A] We thank the reviewer for this suggestion. We have now summarised the mutational\noutcomes of our 16 humanized lines during MAL and AEL by adding Figure 6. An extended\nversion of the experimental evolution outcome can also be found in Supplementary Figure 1.\nMinor Comments\n9) The four differentially expressed proteins in the humanized ancestor at 0 SCBs are referred to in\nTable S11, but should be Table S12.\n[R&A] We apologize for this typo. We revised the numeration of the supplementary tables to reflect\nthe order in which they are cited in the manuscript. Tables S11 and S12 are now Tables S8 and S9.\nWe now cite the correct table (S9) on page 5, line 132.\n10) In line 166, the authors approximate the length of the AEL to ~400 generations. However, in the\nmethods, they report 31 1:100 dilutions. This should correspond to 6.6 generations per transfer,\nresulting in ~200 generations. Is this a typo, or does the difference derive from a different estimate\nof the generations, and if so, which one?\n[R] We tried to calculate the total number of generations based on growth rates before and after\nAEL but we agree with the reviewer that it is more correct to report the theoretical number of\ngenerations based on dilution factors and number of transfers, although this implies no cell death\nand equal fitness across the AEL.\n[A] We revised our calculations and used the formula n=log2(DF) to estimate the total number of\ngenerations for AEL, where DF=dilution factor (1:100). This gives us an estimate of 6.64\ngenerations per serial transfer, and 206 total generations for the AEL, in agreement with the\nreviewer's calculations. We replaced the old value on page 8, line 172 and added the explanation for\nthis calculation in the methods (page 33, lines 802-805).\n11) In the text and figures, serial transfers are referred to as ‘st’ (line 166); however, to avoid\nconfusion, I believe they should be referred to with capital letters, as for the SCBs (ST for singular,\nSTs for plural, as present in the methods).\n[R&A] We changed \"scb\" to \"SCBs\" and \"st\" to \"STs\" across the text, figures and tables.\nReviewer #3 (Remarks to the Author):\nThis study integrates mutation accumulation (MAL) with adaptive evolution (AEL) to chart the\nadaptive molecular landscape triggered by telomeric sequence reprogramming. The authors\ndocument early genome instability, proteome-wide reprogramming, and fitness decline, followed by\nrecovery via two recurrent routes: (i) increased TBF1 dosage—often through chrXVI aneuploidy\nand finer structural variants—and (ii) attenuating mutations in the DNA damage response (DDR).\nThe work addresses a fundamental question—how telomere sequence changes drive long-term\norganismal adaptation—with a coherent multi-omics framework and notable novelty. The findings\nare of high conceptual interest and merit publication after additional validation.\nMajor concerns:\n12) The data presented in Figure 2E, which shows the proportion of T2AG3 sequences across\nchromosomes, is particularly interesting. We noted some variability in the humanization efficiency\namong different clones and chromosomes, an observation that could be further highlighted and\ndiscussed in the main text. To enhance the impact of this work, it would be helpful if the authors\ncould elaborate on the potential biological or technical reasons behind the observed heterogeneity.\nAdditionally, some discussion on the challenges of achieving complete humanization across an\nentire chromosome would be very insightful for the reader community.\n[R] We thank the reviewer for this suggestion. Telomerase does not act at all telomeres at every cell\ncycle, but the probability of a telomere to be elongated increases as its length decreases. Moreover,\nthe number of nucleotides added by telomerase is not always the same. When a telomere\napproaches critical length, telomerase will elongate it to restore a normal length (Teixeira et al.,\n2004; Wellinger & Zakian, 2012). In the case of humanized yeasts, this implies that a minimum\ntract of TG repeats will never get eroded and replaced by T AG repeats.\n1-3 2 3\nHumanized yeasts' telomeres are subject to a higher turnover, as shown in previous studies (Bah et\nal., 2011). This, together with the stochasticity of telomerase's action in terms of number of\nnucleotides added and probability of action at each single chromosome-end during the cell cycle,\nexplains a great part of the heterogeneity that we observed.\nThe high turnover of humanized yeasts' telomeres also implies that they can easily reach critical\nlength and activate DDR. This is compensated by the abundant amplifications of subtelomeric ITS\nand Y' elements that are reminiscent of type I survivors, which usually have short telomeres. Yeasts\nwith telomere defects can also survive through minor amplifications of subtelomeric elements\ncoupled with extremely long telomeric repeats, that are maintained through rolling circle replication\n(type II survivors) (Wellinger & Zakian, 2012). The fully humanized telomere in Hum15 TELIVR\n(>3 kb) is consistent with a type II survivor.\nOverall, the maintenance of an internal TG tract, as well as the global heterogeneity in telomere\n1-3\nlength, can be explained by the combined action of telomerase dynamics, its regulation by telomere\nlength and alternative-lengthening-of-telomeres mechanisms.\n[A] We further discussed the heterogeneity and dynamics of humanized yeasts' telomeres in the\nresults section (pages 8-9, lines 191-198 and 212-217).\n13) There are certain differences between human and yeast telomere-associated proteins. Would\noverexpressing some human telomere proteins in yeast make the study of humanized telomeres in\nyeast more meaningful?\n[R&A] Previous studies have shown that the human telomerase RNA and the telomerase catalytic\nsubunit (TERT) can be successfully expressed in yeast, they localize to the nucleus and show\nprocessivity in yeast cell extracts. However, no telomerase activity could be detected at yeast\ntelomeres in vivo, meaning that it might be necessary to express additional human-specific\ncomponents in order to reconstitute a functional human telomerase holoenzyme in yeast (Bah et al.,\n2004). To our knowledge, no studies have tried to express human telomere binding proteins (TRF1,\nTRF2) together with telomerase components in wild type or humanized yeasts. Given the high\nspecificity of TRF1/TRF2, it is unlikely that their expression in wild type yeasts would lead to a\nfunctional complementation of their yeast counterparts. In humanized yeasts, the telomeric DNA is\nbound by Tbf1p. The simultaneous presence of Tbf1p with TRF1 and TRF2 would lead to\ncompetition for the same binding sites. The resulting telosome structure will be ultimately dictated\nby which proteins have the strongest affinity for the T AG repeats.\n2 3\nTelomere biology is interconnected with DNA repair pathways, cell cycle regulation, mitochondrial\nbiology and other core cellular functions. A change in telomere binding proteins would disrupt these\ninterconnected networks leading to unpredictable fitness outcomes. Future studies expressing\nadditional components of the telomerase holoenzyme and the shelterin complex will help to answer\nthese questions and test the plasticity of telomere functions.\n14) The authors employed two laboratory evolution protocols—mutation accumulation lines (MAL)\nand adaptive evolution lines (AEL)—to effectively investigate the evolution of telomere sequences.\nIt is suggested that the authors clarify in the Discussion section the distinct mutational outcomes\nassociated with each evolution method and provide a rationale for these differences. For instance,\nwhile the authors observed chromosome XVI aneuploidy specifically in the MAL group, other\nstudies have indicated that aneuploidy adaptations are more commonly associated with adaptive\nevolution lines.\n[R] The aneuploidies in MAL are less expected given the close to neutral scenario. However, under\nstrong selection and given their relatively high occurrence rate, it is reasonable to observe them in\nMAL despite their relatively small population size. This has been observed also in MAL performed\nunder rapamycin selection with repeated chromosome XII amplifications to counteract the loss of\nrDNA (Li et al., 2023).\n[A] We clarified this interpretation in the discussion section (page 27, lines 648-653).\nMinor concerns:\n15) The key conclusions rely on assembly and quantification of terminal repeats using long reads. It\nis recommended that the Methods section be written in more detail.\n[R] We annotated TG and T AG repeats in the genome assemblies by using a custom Perl script\n1-3 2 3\nadapted from our recently published pipeline YeaISTY (D’Angiolo et al., 2023). This script inspects\nthe scaffolds of genome assemblies and detects single units of telomeric motifs (CA, CCA, CCCA,\nTG, TGG, TGGG). If units are consecutive to one another they are considered as part of the same\ntelomeric repeats stretch and its start/end points are assigned, respectively, as the position of the\nfirst base of the first unit and the position of the last base of the last unit. Once we obtained a list of\ntelomeric repeats stretches and their start/end points in the scaffolds, we merged the repeats which\nwere interspaced by only 1 bp into a unique stretch by using the software mergeBed included in the\nsuite bedtools. Then, we filtered the list to keep only the stretches longer than 10 bp and we\nmanually inspected them to eliminate breaks that were likely due to sequencing errors. The length\nof the telomeric repeats stretch was then calculated as end-start + 1. The interstitial stretches were\nannotated as ITS if they closely preceded a Y' element, while the stretches at the termini were\nannotated as telomeres.\n[A] We split the methods sections regarding annotations of subtelomeric elements (X and Y') and\nannotations of telomeres and ITS, which were originally merged in a single paragraph. We added a\ndetailed description of the approach we used to annotate telomeres and ITS from long-read genome\nassemblies (page 42, lines 1096-1110).\n16) The key conclusions rely on changes in doubling time; the Results section should add the\ncalculation methods, and explain why correlations were computed on the log scale or not log scale.\n[R] In order to get a comprehensive view of how telomere humanization impacts yeast growth, we\nused two complementary approaches. First, we measured growth in the same medium used for our\nMAL and AEL (Leucin/Tryptophan dropout) by monitoring the optical density in liquid medium\nover a 72 hours timespan in a TECAN plate reader. We derived the doubling time from the resulting\ndata by using the software PRECOG and averaging the values of three biological replicates per\nsample. In PRECOG, the doubling time is calculated as the mean of the five highest slopes in the\nexponential growth phase (Fernandez-Ricaud et al., 2016).\nSecond, we asked whether the impact of telomere humanization would be different in other\nconditions that were not used in our experimental evolution. Given the high number of conditions\ntested we used the high-throughput and automated microbial phenomics platform Scan-o-matic,\nwhich monitors colony sizes on agar plates using desktop scanners and derives growth curves from\nthese images. In Scan-o-matic, the doubling time is calculated as the maximum slope of the growth\ncurve, using five timepoints for a local regression (Zackrisson et al., 2016).\nThe impact of telomere humanization on growth combined with the impact of multiple conditions\ncan result in very different doubling time values across the conditions and samples, making it hard\nto appreciate the effect of each environment in a single plot. To improve visualization and clarity,\nwe expressed the absolute values as relative doubling time respect to a common reference strain\n(wild type ancestor at 2 SCBs) and we converted them to a logarithmic scale.\n[A] We clarified in the results section that we used two different approaches to measure growth and\nwe briefly explained them (page 12, lines 259-269). We included more details on how we calculated\nthe doubling time in the two approaches and we justified our choice of showing part of the growth\ndata in logarithmic scale in the caption of Figure 3. We improved the clarity of the relative methods\nsections and we changed the title from \"Estimation of the growth performance in liquid medium\" to\n\"Estimation of the growth performance in Leucine/Tryptophan dropout\" to better clarify that this\nparagraph refers to the first approach.\n17) The combination of mutation accumulation (MAL) and adaptive evolution (AEL) protocols is a\npowerful design that effectively distinguishes stochastic mutations from adaptive ones. However,\nthe rationale for selecting the specific endpoints of 100 single-cell bottlenecks for MAL and 31\nserial transfers for AEL is not explicitly justified. It would be important to clarify whether these\nthresholds were defined based on preliminary biological observations or statistical considerations.\n[R] We made different considerations to plan the duration of our MAL and AEL, and these are\nexplained below:\nMAL - One of the goals of our study was to characterize the long-term impact of telomere\nhumanization on organismal fitness, including growth, lifespan and mutation rate. To plan the\nduration of our MAL, we considered S. cerevisiae's mutation rate and the number of generations\nneeded to substitute yeast telomeric repeats with humanized ones. Previous studies showed that the\nreplacement of TG with T AG repeats is gradual and only reaches a steady state after 1200\n1-3 2 3\ngenerations (Bah et al., 2004), hence we wanted our MAL to go beyond that number of generations.\nClassical approaches to measure mutation rates in microbes use mutation accumulation lines (MAL)\nin which cells are propagated through SCBs to minimize selection. Since S. cerevisiae has very low\nbase substitution rates (10-9 -10-10/bp/generation) (Barrick & Lenski, 2013; Sharp et al., 2018; Tattini\net al., 2019) it is essential to propagate cells for a long time to obtain enough mutations for\nstatistical comparisons. Thus, we initially planned our MAL to last for at least 2000 generations.\nAssuming that yeasts perform ~20 generations at every bottleneck, our minimal MAL duration was\nAEL - The duration of AEL experiments is usually between few hundreds to a thousand generations\n(LaBar et al., 2020). Previous AEL carried out in our laboratory consisted of ~15 serial transfers,\nthus that number was taken as our minimum baseline (Li et al., 2023; Vázquez-García et al., 2017).\nSince the fitness defect caused by humanized telomeres was very pronounced, we doubled the\nnumber of serial transfers to 30 to have a higher chance of recovering adapted lines. At 31 STs, we\nmeasured the growth rate of our evolved lines and their ancestors and we found that our humanized\nlines had improved their growth, with some of them reaching the same growth rate as the wild\ntypes. Thus, we stopped AEL at that point.\n[A] We clarified in the methods how the thresholds of 100 SCBs and 31 STs were chosen (pages 32-\n33, lines 768-772 and 798-802).\n18) Line 257 mentions that although yeast mitosis has been restored during evolution, chronological\nlifespan has not been restored. Mechanism behind this could be further discussed.\n[R] In both AEL and MAL, we let the cells grow for either 2 or 3 days, transferred them to another\nagar plate or flask and let them grow again. During these protocols, the maximum timespan that\nyeasts spent between cycles was 3 days. After 3 days, both humanized and wild type yeasts had\nreached stationary phase but their viability had not started to decrease yet. Thus, our transfer regime\nimposes that growth rate is constantly under selection during AEL, while there is no selection on\nchronological lifespan.\nPrevious literature has shown that humanized telomeres tend to be shorter and are subject to higher\nturnover than native ones. Additionally, humanized yeasts cannot grow without telomerase (Bah et\nal., 2011). In this context, an increase in growth rate caused by the inactivation of cell cycle\ncheckpoints (either through point mutations in DDR genes or TBF1 amplification and inhibition of\nDDR) might further exacerbate the replication stress and impose a constraint on lifespan due to\nlimiting telomere length, higher genome instability or other yet unknown factors elicited by\ntelomere dysfunction. This is in line with the observation that humanized yeasts promptly improve\ntheir growth but also shorten their lifespan upon the occurrence of chrXVI aneuploidy or point\nmutations in DDR components (Figures 4d and 5b, Supplementary Figures 4a and 7e).\n[A] We now state in the results section (page 12, line 280) that our evolutionary regimes do not\nimpose any selection on chronological lifespan, explaining why this shortens upon MAL and AEL.\nMoreover, we discussed potential explanations for the tradeoff between growth and lifespan that we\nobserved upon DDR mutations and TBF1 amplification (page 27, lines 628-634).\n19) The discussion could be strengthened by connecting telomere sequence evolution with the\nbroader concept of cross-species telomere complex compatibility. Since telomere maintenance\nrelies on co-evolution between telomeric DNA and binding proteins, contrasting the yeast\nhumanization model with examples from other taxa would help contextualize the evolutionary\nadaptability and constraints observed here.\n[R] We thank the reviewer for this suggestion. The concept of cross-species telomere complex\ncompatibility can be viewed in terms of the possibility to switch telomeric sequences between\nspecies or express a functional telomerase across species.\nYeasts can tolerate modifications in their telomeric repeats: S. cerevisiae strains with mutations in\ntheir telomerase RNA template were generated by (Lin et al., 2004) although they showed growth\ndefects and telomere length dysregulation. S. cerevisiae yeasts with humanized telomeres are viable\nalthough they have fitness defects (Henning et al., 1998). More recently, (Červenák et al., 2025)\nreplaced the telomeric repeats of Yarrowia lipolytica with those from human, Yarrowia\nyakushimensis or Candida hispaniensis and showed that the new telomeric repeats were readily\naccumulated at its telomeres. A variant telomeric repeat has been identified also in a human family\n(Hinchie et al., 2024).\nThe telomerase catalytic subunit (TERT) and its associated RNA are sufficient for telomere\nsynthesis in vitro. Previous studies have examined the compatibility between these two components\nacross species. The probability of functional complementation is higher when taxa are\nphylogenetically close. For example, telomerase RNAs can be successfully swapped between the\nciliates Glaucoma chattoni and Tetrahymena thermophila, and the rabbit telomerase RNA is\ncompatible with the human TERT (Bhattacharyya & Blackburn, 1997; Xiang et al., 2000). Instead,\nthere is only partial complementation between mouse and human: while the mouse telomerase RNA\ncannot function with human TERT, the human telomerase RNA can function with mouse TERT\n(Beattie et al., 1998; Boklan et al., 2002; Martín-Rivera et al., 1998). A partial complementation is\nobserved and between yeast and human: in fact, the human telomerase RNA and TERT can be\nsuccessfully expressed in yeast, they localize to the nucleus and show processivity in yeast cell\nextracts. However, no telomerase activity could be detected at yeast telomeres in vivo (Bah et al.,\n2004).\n[A] We strengthened the discussion by describing the cross-species telomere complex compatibility\nin terms of the possibility to switch telomeric sequences between species or express a functional\ntelomerase across species (page 28, lines 663-685).\n20) Petite mutants in wild-type MALs are discarded without quantifying their incidence in\nhumanized lines or exploring telomere–mitochondria interactions.\n[R] We thank the reviewer for this suggestion and we agree that exploring telomere–mitochondria\ninteractions is exceptionally interesting. Telomeres and mitochondrial biology are closely connected\nand telomere dysfunction is known to affect mitochondrial metabolism through the transcriptional\ndysregulation of mitochondrial biogenesis pathways (Finkel, 2011; Robin et al., 2020; Sahin et al.,\n2011). We carried out initial experiments on our MAL to investigate the impact of telomere\nhumanization on mitochondrial functions. Of note, we found a higher petite formation rate in\nhumanized than in wild type yeasts, that seems in contrast with what occurred in our MAL. It is\npossible that by stochasticity we did not pick any humanized petite while we picked multiple wild\ntype petite. Another likely explanation is that petite colonies disappear in humanized yeasts,\nespecially at later timepoints, due to the fact that humanized yeasts already have growth defects and\nthe lack of mitochondrial DNA might simply be lethal in this background. The observation that\npetite rate decreases from 60 SCBs in humanized yeasts while it remains constant for wild types\nseems in line with this explanation. However, we cannot exclude that the petite rescue might be due\nto chromosome XVI aneuploidy, as it happened with the rescue of mitotic growth.\nWe plan to expand the scope of these experiments and gain a comprehensive view of the interplay\nbetween humanized telomeres and mitochondria in a follow-up work. Our preliminary results are\nstill unpublished but we make them available in this response letter.\n[A] We investigated whether telomere humanization had an effect on mitochondrial biology by\nmeasuring several mitochondrial parameters. First, we tested whether telomere humanization\ninfluenced the rate of petite colony formation (Lipinski et al., 2010). Indeed, humanized yeasts\ngenerated a higher percentage of petite colonies than wild types when plated on YPG medium. This\neffect was already present at 20 SCBs but improved at later timepoints in MAL (Figure a).\nSecond, we estimated the mitochondrial DNA copy number by analysing coverage along three\nmitochondrially-encoded genes: COX2, COX3 and ATP6, as previously described (De Chiara et al.,\n2020). While starting from similar copy numbers at 2 SCBs (~2), humanized yeasts increased their\nmtDNA copies during MAL to a final average of ~3 copies, whereas wild types did not change\n(two-tailed Wilcoxon test, p=0.0002 at 100 SCBs) (Figure b).\nThird, mitochondrial dysfunction is usually characterized by an increase of reactive oxygen species\n(ROS) due to malfunctioning of the electron transport chain. We measured the amount of\nsuperoxide anion (O2•−) in humanized and wild type lines, by staining with dihydroethidium\nfollowed by flow cytometry. We found a clear difference in the amount of superoxide between the\ntwo genetic backgrounds, with humanized yeasts having significantly higher superoxide than wild\ntypes (two-tailed Wilcoxon test, p=0.04) (Figure c).\nFinally, we measured mitochondrial activity and mass by double staining with Mitotracker Red and\nMitotracker Green, respectively, followed by flow cytometry. Humanized yeasts had lower\nmitochondrial activity (two-tailed Wilcoxon test, p=0.007), but higher mitochondrial mass (two-\ntailed Wilcoxon test, p=1.5e-5) than wild types (Figures d-e). These apparently contrasting results\nmay be explained by the fact that replication of dysfunctional mitochondria results in higher\nmitochondrial mass but does not improve their activity. These results are also in agreement with our\nestimation of mtDNA copy number from whole-genome sequencing and with an increase in ROS\ndue to respiration defects. Overall, these experiments show that telomere humanization causes\nmitochondrial dysfunction, and the increase in mitochondrial DNA and mass is likely a\ncompensation mechanism to cope with a defective respiration.\n[Figure Redacted]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have adequately addressed my comments and revised the manuscript accordingly. The final version of the paper has improved substantially following the suggestions of all reviewers. Overall, I am satisfied with the authors’ responses and recommend the manuscript for publication.", "reviewer_2": "The authors have addressed all of my comments and have made substantial efforts to improve the clarity of the numerous experimental methods employed. This work represents a significant contribution to our understanding of telomere plasticity and evolution, and the manuscript is likely to become a reference in the field.", "reviewer_3": "The authors have satisfactorily addressed my concerns. I recommend acceptance."}, "author_response": "[R] Thank you."}]

Decision letter