Mutational landscape of normal epithelial cells in Lynch Syndrome patients

Lynch Syndrome (LS) is an autosomal dominant disease conferring a high risk of colorectal cancer due to germline heterozygous mutations in a DNA mismatch repair (MMR) gene. Although cancers in LS patients show elevated somatic mutation burdens, information on mutation rates in normal tissues and understanding of the trajectory from normal to cancer cell is limited. Here we whole genome sequence 152 crypts from normal and neoplastic epithelial tissues from 10 LS patients. In normal tissues the repertoire of mutational processes and mutation rates is similar to that found in wild type individuals. A morphologically normal colonic crypt with an increased mutation burden and MMR deficiency-associated mutational signatures is identified, which may represent a very early stage of LS pathogenesis. Phylogenetic trees of tumour crypts indicate that the most recent ancestor cell of each tumour is already MMR deficient and has experienced multiple cycles of clonal evolution. This study demonstrates the genomic stability of epithelial cells with heterozygous germline MMR gene mutations and highlights important differences in the pathogenesis of LS from other colorectal cancer predisposition syndromes.

Lynch Syndrome (LS) is the most common cause of hereditary colorectal cancer (CRC), accounting for ~3% of all CRC cases¹. LS patients carrying a germline mutation in one of the MMR genes, typically MLH1, MSH2, or EPCAM deletion, have a lifetime CRC risk of 41–80% and endometrial cancer risk of 40–60%, with median age of onset in the 40s^2,3,4,5. Cancer onset is thought to be a result of inactivation of the remaining functional MMR gene allele, which leads to malfunction of the MMR system. Failure to correct mismatched bases and errors from DNA slippage during replication leads to hypermutation and microsatellite instability (MSI), increasing the chance of acquiring cancer driver mutations and thus the development of adenomas and cancers.

A recent study of individuals with germline POLE or POLD1 exonuclease domain mutations and predisposition to colorectal neoplasia showed that somatic mutation rates were elevated in all cells in all normal tissues examined, with the increased burden being attributed to distinctive mutational signatures⁶. Both POL and MMR proteins are involved in DNA repair during replication, and their loss of function can lead to hypermutation in tumours⁷. However, whether normal tissues in LS display any generalized increase in somatic mutation rate is uncertain^8,9 as is the frequency and timing of further elevations in mutation rates, including those associated with wild-type allele loss or neoplastic change.

In this study, we show that the mutation rates of normal epithelium cells in LS patients are similar to wild-type individuals. We identify a morphologically normal crypt displaying increased mutation burden and mutational signatures consistent with MMR deficiency, potentially representing the earliest stage in LS tumourigenesis. Phylogenetic tree reconstructions demonstrate that the most recent ancestor crypt in LS cancers has accumulated many mutations, including key cancer drivers, before the final clonal expansion. These findings demonstrate the genomic stability of normal tissues in LS patients, whose mutation accumulation patterns during tumour development highlight the unique differences of LS tumourigenesis from other CRC predisposition syndromes.

Mutation rates of normal LS epithelium are not elevated

10 LS patients aged 23–78 were recruited to the study. From normal tissues, intestinal crypts (93 colorectal, 14 small intestine) were laser capture microdissected from eight individuals, endometrial glands from one individual, and gastric glands from one individual. Microscopically completely normal crypts were randomly selected for dissection (“survey crypts”) (Fig. 1a). However, given the theoretical possibility that just a subset of non-neoplastic crypts might have become MMR deficient, an additional series was selected on the basis of specific morphologies including budding, branching, and unusual shapes. Furthermore, we immunohistochemically stained normal tissue from 9 LS patients with antibodies to MLH1, MSH2, or PMS2 to find individual crypts which were negative and which may, therefore, have already lost DNA MMR functions. Crypts were also sampled from three colonic adenocarcinomas from three patients, with one including an adenomatous component at the tumour edge. A total of 132 non-neoplastic crypts/glands including 107 intestinal crypts, 8 endometrial glands, 17 gastric glands, and 20 colorectal neoplasm crypts were individually microdissected and whole genome sequenced to ~33X coverage (Supplementary Fig. 1 and Supplementary Data 1).

a Representative images of H&E and IHC staining of crypts with different morphologies. Except for the MMR deficient crypt (indicated by white arrows), images of H&E and IHC did not represent the same crypt. MMR protein antibodies were used according to the germline MMR gene mutation in respective patients. MMR-d: MMR-deficient; IM: intestinal metaplasia; AdenoCA: adenocarcinoma. Scale bar: 100 μm (i, ii, v, vi, vii, viii, ix, x, xvi, xviii); 50 μm (iii, iv, xi, xii, xiii, xiv, xv, xvii). b–d Boxplots showing the 1.5X interquartile range (whiskers), the first and third quartile (bounds of box), and the median (centre)of the data. Each data point represented a crypt. Two-sided Wilcoxon test: ns = non-significant. b Single base substitutions (SBS) and insertions and deletions (ID) burden of survey crypts from the stomach (n = 5, red), the small intestine (n = 14, blue), the colon (n = 37, green) and the endometrium (n = 8, purple). c SBS and ID burden in different types of colonic crypts (Survey (n = 37), Specific morphologies (n = 50), MMR-d (n = 1), Tumour edge (n = 5)). Crypts with and without driver mutations are indicated in pink and blue, respectively. d SBS and ID burden in non-neoplastic crypts with (n = 28, pink) and without (n = 104, blue) cancer driver mutations. e A phylogenetic tree showing the clonal relationship between non-neoplastic crypts in PD46179. Branch lengths correspond to the number of SBS mutations. SBS mutational signatures are mapped onto tree branches. Each crypt is annotated with its identifier, crypt type, and tissue origin.

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To assess if the microdissected crypts each contained cells originating from a single adult stem cell, the variant allele frequency (VAF) distribution was assessed for each crypt. All crypts showed a VAF peak around 0.5 and a median VAF > 0.3, confirming their clonal nature (Supplementary Fig. 2). The median base substitution mutation rates in morphologically normal crypts/glands (i.e., “survey” crypts) from the stomach, the small intestine, the colon, and the endometrium were 38.5, 72.6, 55.2, and 35.9 substitutions per year, respectively, while the median single base insertion and deletion (ID) rates were 1.6, 2.8, 2.6 and 2.1 per year, respectively (Fig. 1b). Colonic crypts with specific morphologies, including budding, branching, and unusual shapes, and also normal crypts adjacent to the edge of a tumour, showed comparable SBS and ID rates to survey crypts, and the presence of driver mutations was not associated with a particular crypt type (Fig. 1c,   d). To test if the mutation rate of normal colonic crypts from LS patients is different from that of wild type individuals, we combined the LS dataset with data from 424 crypts from 42 individuals sequenced as part of our earlier study of the mutation landscape of the normal colon (Supplementary Data 2)^10,11. We used a linear mixed-effect model to test for differences in the mutation rate between LS and wild type crypts. Likelihood ratio tests showed that including a fixed-effect variable for LS status did not improve the fit of the model (SBS: p = 0.11, ID: p = 0.14), suggesting that if there are differences between the mutation rate of wild type and LS crypts, they are too subtle to be detected in this study. There were, however, specific groups of crypts with modestly elevated mutation burdens, including those from the small intestine of PD43853, who had received prior chemotherapy treatment, and antral mucosal glands of PD45540 who had prior Helicobacter pylori infection and chemotherapy treatment (Supplementary Data 3 and Supplementary Data 4).

To confirm that the mutation rate of normal LS tissue is not generally elevated, clonal organoid cultures were established from normal gastrointestinal adult stem cells (ASCs) from two patients (PD45539 and PD45540) and whole genome sequenced (Supplementary Fig. 3a). Mutation rates over the lifetimes of these patients and mutational signatures were comparable to those of organoids from a wild type individual (Supplementary Fig. 3b, c and Supplementary Data 5). Furthermore, extended culture of LS organoids for >4 months did not reveal an increased in vitro mutation rate compared to organoids from the wild-type individual, confirming the stability of the genome in normal cells from LS patients (Supplementary Fig. 3d and Supplementary Data 6).

A morphologically normal crypt shows early MMR deficiency

We examined intestinal, endometrial, and gastric epithelium from 9 LS patients using immunohistochemistry for MLH1, MSH2, or PMS2 proteins. A single colonic crypt (lo0054) was found to be negative for MSH2 staining in a young patient (PD46179) with a germline MSH2 mutation. Subsequent microdissection of this crypt and whole genome sequencing showed elevated SBS and ID rates compared to survey crypts, with the mutation rate estimated to be 197 SBSs and 37 IDs per year (Fig. 1c). The elevated mutation rates were primarily due to increased rates of SBS1 and SBS44 together with ID1 and ID2 (Fig. 1e and Supplementary Fig. 4d). It is conceivable that the elevated rate of SBS1 (a mutational signature predominantly characterized by C > T mutations at CG dinucleotides) was due to defective processing of G:T mismatches (a known function of the MMR system), and SBS44 is a signature known to be associated with defective DNA MMR system. The crypt harboured two potential driver mutations, NF2 R172fs and PIK3R1 L30fs. Taken together, the evidence indicates that this morphologically normal crypt already had defective DNA MMR function and thus may represent an extremely early stage of the pathogenesis of LS.

Interestingly, and notwithstanding a careful search, this crypt did not show a somatic MSH2 mutation nor an LOH event that would cause inactivation of the wild type MSH2 allele. The absence of an MSH2second hit, despite the elevated mutation burden and evidence of MSI, suggests that some degree of loss of MMR function and emergence of the MSI phenotype can sometimes precede inactivation of the wild type MMR gene allele in LS. The nature of the somatic event that has conferred MMR deficiency in this normal crypt is unclear. However, the observation may suggest a multistep model of progression to full MMR deficiency in LS in which normal crypts first acquire a degree of MMR deficiency, through a currently unknown mechanism, which results in an elevated SBS and ID mutation rate. In turn, this causes mutation and inactivation of the inherited, wild type MMR allele conferring a greater degree of MMR deficiency and an even higher mutation rate that leads to neoplastic changes.

Driver mutations in normal LS epithelium

Of the 132 glands/crypts (intestinal, gastric, and endometrial glands/crypts included) from normal epithelium, 28 (21.2%) had putative cancer driver mutations. Among these 28 glands, 23 (17.4%) had one driver mutation and five (3.8%) had two (Fig. 2a). Altogether there were 30 unique driver mutation events involving 24 cancer driver genes and all were heterozygous (Fig. 2b and Supplementary Data 7). Most of the cancer drivers observed in the gastrointestinal tract involved tumour suppressor genes. APC, the most common cancer gene mutated in CRCs, was mutated in two crypts, one (APC T1556fs) from the duodenum and the other (APC Q8X) from the caecum (Fig. 3a, b). There was no evidence of dysplastic change or difference in mutation burden in crypts carrying driver mutations. Mutations of oncogenes, including KRAS, PIK3CA, and ERBB2, were observed in endometrial glands, again without obvious change in morphology or mutation rate, as previously reported in wild type individuals¹² (Fig. 3c). Four copy number variants (CNVs) and 20 structural variants (SVs) were identified, affecting 15.2% of crypts (Fig. 2c), similar in frequency to wild-type individuals. There was a trend for an increasing number of CNVs/SVs with age (Fig. 2d). No homozygous deletions were observed, and the break points of structural variants did not intersect with any cancer driver genes. Interestingly, a chromosome 3p uniparental disomy (UPD) event was found in an appendix crypt (lo0002) in PD45539, a patient with a germline MLH1 N38K mutation (Fig. 3b). This UPD event, however, reverted the germline MLH1mutation back to wild type, re-endowing the crypt with two functional copies of MLH1.

a Distribution of non-neoplastic crypts with driver mutations and copy number or structural variants. b Driver mutations in non-neoplastic crypts. Asterisks denote frameshift mutations at microsatellite regions (repeat length ≥ 5). c Copy number variants and structural variants in non-neoplastic crypts. Cn-LOH: copy number neutral loss of heterozygosity. d Percentage of non-neoplastic crypts with copy number or structural variants in each LS patient.

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Phylogenetic trees showing the clonal relationship between non-neoplastic crypts in 7 patients (a, b, e, f, g intestinal; c endometrial; d gastric). Branch lengths correspond to the number of SBS mutations. SBS mutational signatures are mapped onto tree branches. Each crypt is annotated with its identifier, crypt type, and tissue origin. Cancer driver mutations and copy number variants are labelled on the corresponding tree branch. Frameshift mutations at microsatellite regions (repeat length ≥ 5) are indicated in bold. Activating mutations of oncogenes are indicated in red. Inactivating mutations of tumour suppressors are indicated in black. c KRAS G12V was an early event and thus annotated on the common branch (red arrow) of the 4 crypts harbouring this mutation (dotted red arrows).

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Mutational processes in the LS epithelium

To investigate the genomic characteristics and mutational processes operating in the crypts of LS patients, phylogenetic trees of crypts were constructed, followed by mutational signature extraction for each patient. Mutational processes in normal epithelium from LS patients were similar to those reported in wild type individuals^10,12. All crypts/glands from LS patients predominantly displayed the clock-like signatures SBS1 and SBS5, with SBS2, SBS13, SBS17b, SBS18, SBS35, and SBS88 also present in some crypts/glands from some patients (Fig. 3). SBS35, a signature associated with platinum-based drug usage¹³, was identified in every intestinal crypt in PD43853 who had undergone a combination of 5-fluorouracil (5-FU), leucovorin, and oxaliplatin (FOLFOX) treatment before surgery (Fig. 3a). SBS2 and SBS13, two mutational signatures associated with APOBEC enzyme activities^14,15, were observed in some duodenal and ileal crypts from PD43853, potentially reflecting the presence of localized DNA editing by cytidine deaminases in the small intestine (Fig. 3a). SBS17b, a sub-signature of SBS17 that has in some instances been linked to 5-FU exposure and oxidative stress environment¹⁶, was observed in most crypts in the pyloric antrum but not the body of the stomach in PD45540 who had been treated with 5-FU (Fig. 3d). In a family with a germline EPCAM deletion, SBS88, a signature associated with colibactin exposure^10,17, was found in every crypt from two siblings (PD46176 and PD46177) in their early twenties, but not in their grandmother (PD46178) (Fig. 3e–g). Colibactin is a genotoxin secreted by pks+ E.coli, typically present in the gut microbiota^18,19. The extensive presence of SBS88 in the two siblings suggests a shared microbiome between them.

We compared the mutation burdens in paired normal and tumour (“adenoma” and “adenoCA”) crypts from three LS patients in whom these tissues were available. In all three patients, the SBS and ID burdens of tumour crypts were ~100-fold and 2000-fold higher respectively than normal crypts sampled at the same time (Fig. 4a). Mutational signature analysis showed that all tumour crypts displayed SBS1, SBS5, previously described DNA MMR deficiency-associated signatures SBS20, SBS21, SBS26, SBS44, and an unknown signature N4^14,20. Phylogenetic tree reconstruction showed that tumour crypts formed a distinct cluster away from normal crypts, and the tumour tree branches arose from a single trunk of at least 50,000 SBSs and multiple driver mutations (Fig. 4b–d). This indicates that many mutations had already accumulated in the most recent common ancestor cell of each tumour after multiple rounds of clonal expansion, with subsequent clonal expansion resulting in the formation of the adenoma or carcinoma. We noticed a greater contribution of SBS1 in tree trunks compared to branches, and MMR deficiency-associated signatures contributed to most of the mutations in branches. However, the exact mechanism underlying such phenomenon is unknown. In PD46175 an adenocarcinoma evolved from a recent ancestor cell that also evolved to generate an adjacent adenoma (Fig. 4c). In all three tumours, somatic wild type allele inactivation of the relevant MMR gene and APC biallelic mutations were present in all tumour crypts and mapped onto the trunk of the tumour cluster, consistent with existing knowledge of one of the tumourigenesis pathways in LS in which adenoma formation requires both MMR deficiency and APC inactivation^21,22.

a Single base substitutions (SBS) and insertions and deletions (ID) mutation density of tumour crypts (blue) compared to survey crypts (black) in the three patients with a paired colon tumour tissue. Two-sided Wilcoxon test: PD46174 (Survey: n = 3; Tumour: n = 6; p = 0.024), PD46175 (Survey: n = 3; Tumour: n = 10; p = 0.007), PD46179 (Survey: n = 4; Tumour: n = 4; p = 0.029) for both SBS and ID. b–d Phylogenetic trees showing the clonal relationship between crypts in the three patients with a paired tumour sample. Branch lengths correspond to the number of SBS mutations. SBS mutational signatures are mapped onto tree branches. Each crypt is annotated with its identifier, crypt type, and tissue origin. Cancer driver mutations and copy number variants are labelled on the corresponding tree branch. Frameshift mutations at microsatellite regions (repeat length ≥ 5) are indicated in bold. Activating mutations of oncogenes are indicated in red. Inactivating mutations of tumour suppressors are indicated in black. For tumour crypts, only the key cancer driver mutations on major tree branches are annotated. Most of the mutations in tumour crypts were frameshift mutations at microsatellite regions, which were consequences of MSI. These mutations were omitted from annotation. The location where each tumour crypt was sampled is shown by the H&E staining of the tumour section to the right of the phylogenetic trees. c For PD46175, adenoma crypts were annotated in blue brackets. d For PD46179, the inserts of the region within the red brackets show crypts that were sampled from another tissue section. These images of crypts were superimposed onto the tumour section to show its relative position. Scale bar of tumour section: 2.5 mm (PD46174 and PD46179); 5 mm (PD46175).

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As many IDs were generated in the tumour crypts, ID mutational signatures were also extracted in the three patients. All tumour crypts showed mutational signatures ID1, ID2, ID7, and an unknown signature N3 (Supplementary Fig. 4a–c). ID1 and ID2, characterized by single insertions and deletions of thymine at poly-T tracts, respectively, are ubiquitous signatures due to polymerase slippage during DNA replication, with increased exposures observed in samples with MMR deficiency^14,20. ID7, on the other hand, is characterized by mono- and di-nucleotide deletions at microsatellite regions and is associated with MMR deficiency^14,20. The unknown signature N3 was characterized by ≥5 bp deletions at sequences with various repeat lengths. Inspection of these deletions revealed that N3 represents polynucleotide deletions at microsatellite regions, presumably due to continual retraction of microsatellites under MMR deficiency (Supplementary Fig. 4e). The morphologically normal, yet MMR deficient crypt (lo0054) of PD46179, showed mutational signatures ID1 and ID2, but not ID7 or N3, suggesting that MMR deficiency was in an early state in which complex IDs had not yet been accumulated in the genome (Supplementary Fig. 4d).

This study presents the genomic landscape of normal epithelial tissues in LS patients. While elevated mutation burdens have been shown in normal tissues from individuals with germline POLE/POLD1exonuclease domain mutations⁶, the same was not observed generally in the epithelium of individuals with germline MMR gene mutations in this study. In fact, the mutation burden of colonic epithelium in LS patients was comparable to that of wild type individuals¹⁰. Mutation of one allele of the POLE/POLD1 genes appears sufficient to perturb the proofreading function of the DNA polymerase and lead to the accumulation of mutations in the genome, whereas mutation of one allele of an MMR gene in the presence of a wild type allele is insufficient to disrupt the MMR system. The protective function of the wild type MMR allele is further illustrated by a recent study of normal tissues from individuals with constitutional MMR deficiency (CMMRD) due to inherited biallelic mutations in MMR genes, in which the mutation rate of normal intestinal epithelial cells ranged from 500 to 3000 mutations per year, 10 to 60-fold higher than that in LS patients²³.

Despite the substantially greater mutation rate in normal tissues from POLE/POLD1 mutation carriers than from MMR gene mutation carriers, the estimated risk of colorectal cancer by age 70 is only moderately higher in POLE (90%) than in MLH1/MSH2 (41–48%) gene mutation carriers^4,24. These observations indicate that the elevated mutation rate present in all normal colorectal epithelial cells in POLE/POLD1 mutation carriers leads to a higher rate of development of adenomas compared to wild-type and LS individuals. However, the risk of conversion from adenoma to carcinoma may be greater in MLH1/MSH2 gene mutation carriers, perhaps due to higher ID mutation rates that supervene once the wild-type MMR gene allele is lost in individual cells.

We identified a single morphologically normal but MMR deficient colonic crypt in a young LS patient with an inherited MSH2 mutation. The presence of such crypts in normal colorectum from LS patients has been previously reported. The MMR deficient crypts, as evidenced by loss of MMR protein staining and demonstration of MSI, were found with an estimated frequency of ~1 MMR deficient crypt focus per cm² of mucosa^25,26. In this study, we were able to analyse the genome of such a crypt demonstrating the emergence of MMR deficiency-associated mutational signatures and elevated numbers of SBS and ID mutations. This crypt, with its distinctive mutational patterns, may therefore represent an example of a rare, dispersed subpopulation of cells in the intestine of LS patients which are in the earliest stages of progression from normal to cancer cell. If so, given the relatively small number of colorectal neoplasms that develop in LS patients, only a very small minority of such cells with MMR deficiency and elevated mutation rates ever become symptomatic neoplasms. Interestingly, despite thorough interrogation of the whole genome for mutations, CNV or SV, no MSH2 second hit could be identified. This suggests the existence of a state in which mutations accumulate in the crypt and MSI starts to emerge before the MMR gene second hit takes place. Indeed, this state may contribute to the emergence of fully defective MMR function by contributing the second hit. The biological mechanism underlying this state is unclear though. Nevertheless, we do not exclude the possibility that the absence of MMR gene second hit was merely due to undetected mutations/ inactivating mechanisms given the analysis pipeline and technology platform used in this study.

Based on the phylogenetic tree reconstruction on tumour crypts, we showed that MMR deficiency and APC inactivation are the earliest events in LS tumour progression, with hotspot mutations in other cancer genes such as those in KRAS and TP53 serving as late cancer drivers. While we could not delineate the sequence of mutations within a trunk of mutations, 2 of the 3 tumours had APC frameshift mutations at microsatellite regions, possibly hinting that APC inactivation came after MMR deficiency in the 2 tumours. Overall, our findings are consistent with the pre-existing tumourigenesis pathways described in LS^21,22.

In conclusion, this study demonstrates that the overwhelming majority of normal epithelial crypts with germline heterozygous MMR gene mutations are genomically stable. In future work, larger scale systematic screening for and genetic profiling of the small subpopulation of MMR deficient crypts in the normal epithelium of LS patients will clarify the earliest molecular events a normal colorectal stem cell requires in order to acquire MMR deficiency and subsequently progress to neoplastic transformation.