Aging and carcinogenesis coincide with the accumulation of DNA damage and mutations in stem and progenitor cells. Molecular mechanisms that influence responses of stem and progenitor cells to DNA damage remain to be delineated. Here, we show that niche positioning and Wnt signaling activity modulate the sensitivity of intestinal stem and progenitor cells (ISPCs) to DNA damage. ISPCs at the crypt bottom with high Wnt/β‐catenin activity are more sensitive to DNA damage compared to ISPCs in position 4 with low Wnt activity. These differences are not induced by differences in cell cycle activity but relate to DNA damage‐dependent activation of Wnt signaling, which in turn amplifies DNA damage checkpoint activation. The study shows that instructed enhancement of Wnt signaling increases radio‐sensitivity of ISPCs, while inhibition of Wnt signaling decreases it. These results provide a proof of concept that cell intrinsic levels of Wnt signaling modulate the sensitivity of ISPCs to DNA damage and heterogeneity in Wnt activation in the stem cell niche contributes to the selection of ISPCs in the context of DNA damage.
New genetic results suggest that DNA damage‐induced modulation of Wnt signaling determines radio‐sensitivity of intestinal stem‐ and progenitor cells.
Niche positioning‐associated Wnt signaling activity modulates sensitivity of intestinal stem and progenitor cells to DNA damage.
Wnt signaling enhances DNA damage responses in intestinal stem and progenitor cells.
Intestinal stem and progenitor cells with low Wnt activity preferentially survive in the context of DNA damage and telomere dysfunction.
Instructed enhancement of Wnt signaling increases radio‐sensitivity of intestinal stem and progenitor cells while inhibition of Wnt decreases it.
There is increasing evidence that the functionality of stem cells in adult tissues decreases during aging (for review see Liu & Rando, 2011; Sperka et al, 2012), which seems to be driven by both age‐dependent alterations in stem cell self‐renewal pathways and the accumulation of molecular damages. An accumulation of DNA damage occurs in various tissues and in stem cells of aging mice and humans (Herbig et al, 2006; Rossi et al, 2007; Jiang et al, 2008; Rube et al, 2011). DNA damage impairs stem cell maintenance and function in adult tissues (Rudolph et al, 1999; Armanios et al, 2007; Rossi et al, 2007; Hoeijmakers, 2009; Sperka et al, 2012; Tumpel & Rudolph, 2012). The accumulation of DNA damage and mutations in stem and progenitor cells also contribute to cancer formation during aging, and it was shown that adult tissue stem cells represent the cell type of origin of cancer formation in mouse models (Barker et al, 2009). Emerging data indicate that adult tissue stem cells accumulate mutations that contribute to cancer initiation during aging (Busque et al, 2012; Jan et al, 2012; Welch et al, 2012; Genovese et al, 2014; Jaiswal et al, 2014; Xie et al, 2014).
Molecular mechanisms that influence checkpoint induction and selection of mutant stem and progenitor cells in the context of DNA damage and aging remain yet to be delineated. It is currently not known whether stem cell self‐renewal pathways influence the induction of DNA damage checkpoints in adult tissue stem and progenitor cells. Canonical Wnt/β‐catenin signaling represents a critical self‐renewal pathway, which is required for the maintenance of stem cells in adult mammalian tissues (Reya & Clevers, 2005; van Amerongen et al, 2012). Inactivation of essential components of canonical Wnt signaling results in defects in the maintenance of intestinal stem cells (Fevr et al, 2007; de Lau et al, 2011) and hematopoietic stem cells (HSCs) (Reya et al, 2003; Fleming et al, 2008; Luis et al, 2009). However, also the over‐activation of canonical Wnt signaling is detrimental for stem cell maintenance and leads to premature exhaustion of HSCs (Scheller et al, 2006; Huang et al, 2009; Lane et al, 2010).
Alterations in Wnt signaling were implicated in aging of adult tissues (Brack et al, 2007; Liu et al, 2007; Ye et al, 2007). Specifically, it was shown that increased activity of canonical Wnt/β‐catenin signaling leads to the dysfunction of muscle stem cells during aging (Brack et al, 2007). In the small intestine, Paneth cells build the stem cell niche and provide key signals such as Wnt ligands for the maintenance of intestinal stem cells. Since Paneth cells exclusively sit at the bottom of the crypts, they create a Wnt gradient in the crypts with highest level at the crypt base and lower levels toward the upper part of the crypts (van der Flier et al, 2009; Itzkovitz et al, 2012). Despite numerous findings on the role of Wnt signaling in controlling the maintenance and plasticity of intestinal stem and progenitor cells (ISPCs), mechanistic studies on the influence of Wnt/β‐catenin signaling on the maintenance of ISPC in the context of DNA damage have not been reported. Here, we provide experimental evidence that niche positioning‐dependent differences in the level of canonical Wnt/β‐catenin signaling modulate the sensitivity of ISPCs to DNA damage in the intestinal epithelium of mice. Cells with high Wnt/β‐catenin activity (Wnthi) at the crypt base are more sensitive to DNA damage‐induced depletion compared to Wntlo border cells at position 4. The increased sensitivity of Wnthi cells to DNA damage is not associated with differences in cell cycle activity but involves a feed‐forward loop of transient Wnt activation in response to DNA damage and amplification of DNA damage checkpoints in cells with high Wnt signaling activity. Of note, instructed activation of canonical Wnt signaling enhances radio‐sensitivity of ISPCs, while inhibition of Wnt signaling decreases it, both in cell culture and in vivo. These findings could be relevant for the accumulation of genetic alterations and the clonal selection of stem and/or progenitor cells in the context of DNA damage during aging and carcinogenesis as well as for therapeutic targeting of cancer stem cells with elevated levels of DNA damage.
Positioning within the niche and levels of LGR5 expression discriminate intestinal ISPCs with different Wnt/β‐catenin signaling activity
Recent studies revealed a high plasticity of ISPCs in the basal crypts of the intestinal epithelium. It was shown in lineage tracing experiments that LGR5‐positive cells at the crypt base in position 1 and 2 represent intestinal stem cells (Barker et al, 2007). In addition, LGR5+ cells in the border region (position 3/4 of the crypt) have true stem cell activity and can replace stem cells at the crypt base (Ritsma et al, 2014). However, it was also shown that LGR5‐negative TA cells can revert to organoid‐forming stem cells in culture when exposed to Wnt3A (Sato et al, 2011) as well as in response to tissue injury in vivo (van Es et al, 2012), but these events are rare and LGR5+ cells were found to be essential for intestinal regeneration and mouse survival in response to irradiation (IR) (Metcalfe et al, 2014). The main aim of this study was to delineate the potential influence of Wnt/β‐catenin signaling on the survival of the total population of ISPCs in response to DNA damage rather than to re‐investigate the discrimination of intestinal stem and progenitor cells and the plasticity of early progenitors to convert into stem cells or vice versa. Therefore, LGR5+ cells in position 1–4 cells are altogether referred to as “stem and progenitor cells (ISPCs)” from here on.
LGR5+ ISPCs are placed in between and adjacent to the Paneth cells in position 1–4 of the crypt base (Barker et al, 2007). Paneth cells produce the Wnt/β‐catenin stimulatory ligand Wnt3, which represents an essential factor for adapting freshly isolated LGR5+ ISPCs to cell culture (Sato et al, 2011). Although Paneth cells are dispensable for the maintenance of intestinal stem cells in vivo (Kim et al, 2012), it was shown that intestinal stem cells that are located in between the Paneth cells at the very bottom of the crypts (also known as “crypt base columnar stem cells—CBCs” located at position 1 and 2 of the crypt base) express higher levels of Wnt target genes such as Lgr5, Ascl2, and Sox9 compared to position 4 cells that are located above the Paneth cells and are therefore also known as “border cells” (van der Flier et al, 2009; Itzkovitz et al, 2012; Ritsma et al, 2014). These previous studies indicated that Paneth cells generate a Wnt gradient‐exposing cells at the crypt base that are surrounded by multiple Paneth cells to a higher Wnt3 level than border cells that are located above the Paneth cells, thus only contacting 1 or 2 Paneth cells. In agreement with previous studies, immunofluorescence staining of LGR5‐GFP expression and Musashi‐1 (Msi1) (both direct targets of Wnt/β‐catenin signaling) confirmed higher Wnt signal activity in position 1–2 cells compared to cells at position 4 (Fig 1A–D). Fluorescence‐activated cell sorting (FACS) of freshly isolated crypts showed that intestinal crypts contained LGR5+ cells with relatively higher (LGR5hi) or lower (LGR5lo) LGR5‐GFP expression (Fig 1E). Aside from the stability of the GFP protein, it was shown that transcriptional control of LGR5 expression, directly targeted by Wnt/β‐catenin signaling, contributes to differences in the level of GFP expression in the population of LGR5‐positive intestinal ISPCs in this reporter strain (Munoz et al, 2012). To determine whether FACS can be used to separate ISPCs with different intrinsic levels of Wnt signaling activity, four subpopulations of cells were FACS‐purified from the intestinal epithelium of the LGR5‐GFP reporter mouse strain based on the level of GFP expression: (i) LGR5hi‐high, (ii) LGR5hi‐low, (iii) LGR5lo‐high, (iv) LGR5lo‐low (Fig 1F). qPCR analysis revealed that the LGR5‐GFP expression level correlated with the expression level of several Wnt target genes including Axin2, Ascl2, and Msi1 (Fig 1G–I), indicating that FACS can be employed to efficiently purify ISPCs with different levels of Wnt signaling. Wnt activity showed an inverse correlation with expression of some differentiation markers (Fabp1, Chga, Dclk1, Supplementary Fig S1). Together, these experiments revealed that FACS purification based on the LGR5 reporter expression separates ISPCs with high or low Wnt signaling activity and immunostaining indicated a correlation between the level of Wnt target gene expression (LGR5 and Msi) and the localization of cells in the basal crypt (position 1/2 > position 4).
Telomere dysfunction associates with suppression of Wnt signaling in ISPCs of aging telomerase‐deficient mice
Homozygous deletion of the RNA component of telomerase (mTerc) accelerates telomere shortening and leads to an age‐dependent accumulation of telomere dysfunction in the third generation of knockout mice (G3 mTerc−/−) compared to wild‐type mice (mTerc+/+). As DNA damage accumulates in G3 mTerc−/− mice, we here used these mice as a model of age‐dependent DNA damage accumulation. Telomere dysfunction in aging G3 mTerc−/− mice induces p53‐dependent impairments of stem cell maintenance and premature aging of somatic tissues (Choudhury et al, 2007; Begus‐Nahrmann et al, 2009; Sperka et al, 2011). The intestinal epithelium is an organ compartment with very high rates of cell turnover, which is strongly affected by aging‐associated accumulation of telomere dysfunction in telomerase‐deficient mice (Choudhury et al, 2007; Schaetzlein et al, 2007; Sperka et al, 2011).
The canonical Wnt signaling pathway has been identified as a major regulator of stem cell maintenance in the intestinal epithelium (Reya & Clevers, 2005). To see whether this pathway is affected in response to telomere dysfunction, gene expression analysis was employed. mRNA expression analysis of components of the canonical Wnt/β‐catenin signaling pathway revealed a significant suppression of various components of the Wnt signaling pathway in freshly isolated basal intestinal crypts of 12‐ to 16‐month‐old G3 mTerc−/− mice compared to age‐matched mTerc+/+ mice (Fig 2A). A significant suppression of Axin2 was also detected in freshly isolated, highly purified, LGR5+ ISPCs from 12‐ to 16‐month‐old G3 mTerc−/− mice compared to age‐matched mTerc+/+ mice (Fig 2B).
Telomere dysfunction was previously shown to repress the ability of LGR5+ ISPCs to grow organoids in cell culture (Sperka et al, 2011). To test whether suppression of Wnt signaling was associated with this deficiency, intestinal organoids were re‐derived from primary cultures after two passages, 18 days after initiating cultures from freshly isolated basal crypts from 2‐month‐old G3 mTerc−/− mice or mTerc+/+ mice. mRNA analysis revealed a significant suppression in the expression of several components of the Wnt pathway, but an increase in the expression of the DNA damage marker p21 in organoids derived from intestinal crypts of G3 mTerc−/− mice compared to mTerc+/+ mice (Fig 2C).
Telomere dysfunction leads to preferential depletion of ISPCs with high Wnt activity
Together, the above data showed that telomere dysfunction and activation of DNA damage signals associate with the suppression of canonical Wnt/β‐catenin signaling in the ISPC compartment. DNA damage accumulates in the intestine of aging G3 mTerc−/− mice, and this leads to impairments in the maintenance of ISPCs and an increasing atrophy of the intestinal epithelium in aging G3 mTerc−/− mice (Begus‐Nahrmann et al, 2009; Sperka et al, 2011). In agreement with these studies, FACS analysis revealed a significant depletion of ISPCs in 12‐ to 16‐month‐old G3 mTerc−/− mice compared to age‐matched mTerc+/+ mice, but not in 2‐ to 3‐month‐old mice (Fig 3A, C and D, Supplementary Fig S2). Interestingly, this age‐dependent decrease in ISPCs was more pronounced in the fraction of LGR5hi cells (Fig 3B and E–G). Moreover, within the LGR5hi cells, the subpopulation of LGR5hi‐high cells was preferentially depleted compared to the subpopulation of LGR5hi‐low cells (Fig 3H–J, see Fig 1F for gating of subpopulations from the total population of LGR5+ cells). Histological analysis indicated that surviving LGR5+ cells in 9‐month‐old G3 mTerc−/− mice exhibited weaker GFP staining intensity and were mostly located at position 4 above the Paneth cells, whereas LGR5+ cells at the crypt base that were located in between the Paneth cells (position 1 and 2) were preferentially depleted (Fig 3K–O).
The use of multiple markers confirmed that telomere dysfunction led to a depletion of ISPCs at position 1–2 of the basal crypt including: (i) morphological markers depicting ISPCs as small spindle‐like cells located adjacent to Paneth cells at the crypt base (Barker et al, 2012) (Fig 4A–C, Supplementary Fig S3), (ii) PCNA immunofluorescence distinguishing proliferative ISPCs from post‐mitotic Paneth cells (Fig 4D–F), (iii) Msi1 immunofluorescence staining for ISPCs at position 1–4 of the basal crypt (Kayahara et al, 2003; Rezza et al, 2010) (Fig 4G–I), and (iv) olfactomedin 4 (Olfm4) in situ hybridization: Olfm4 is a Notch target gene but is not directly regulated by Wnt (van der Flier et al, 2009) and labels ISPCs in position 1–4 of basal crypts (Fig 4J and K).
γ‐irradiation‐induced DNA damage leads to preferential depletion of ISPCs with high Wnt signaling
To determine the DNA damage sensitivity of ISPCs in a different model, we analyzed the number of ISPCs in the intestine of 3‐month‐old mTerc+/+ mice in response to acute exposure to γ‐irradiation. Immunohistochemistry analysis showed a rapid depletion of PCNA‐positive (PCNA+) ISPCs at the crypt base (position 1 and 2 at 24–48 h after IR) but a recovery of these cells at day 4–6 after IR (Fig 5A–G). In contrast, PCNA+ cells located above the Paneth cells (position 4) were maintained after IR (Fig 5A–G). To verify that γ‐irradiation led to the depletion of position 1–2 cells, Wnt‐independent markers (Olfm4, Fig 5H and I) and Wnt‐dependent markers (LGR5 and Msi1, Fig 5J–N, Supplementary Fig S4A–C) were employed. Staining for GFP expression in LGR5‐GFPki mice indicated a depletion of cells at the bottom of the crypt at position 1 and 2 reaching a maximum at 24 h to 2 days after IR (Fig 5J–N), and a preferential maintenance of position 4 cells with weaker GFP staining intensity (Supplementary Fig S4D). At later time points (4 days after IR), a recovery of LGR5+ cells at position 1–2 occurred (Fig 5N). Staining of intestinal crypts for the expression of Olfm4 and Msi1 confirmed the depletion of ISPCs at the crypt bottom at 24 h after IR (Fig 5H and I, Supplementary Fig S4A–C).
To quantify the survival of LGR5lo cells and LGR5hi cells, FACS was used to analyze intestinal crypt cells from irradiated mice at different time points after IR (3–24 h). At 3 h after IR, there was a transient increase of LGR5+ cells indicative of an IR‐mediated induction of Wnt signaling (Fig 5O). However, 24 h after IR, a significant reduction of LGR5+ cells occurred (Fig 5O) predominantly affecting the subpopulation of LGR5hi cells compared to LGR5lo cells (Fig 5P–U). Gating of sub‐populations within the population of LGR5hi cells revealed that the LGR5hi‐high cells were preferentially depleted compared to the LGR5hi‐low cells (Fig 5V–X, see Fig 1F for gating of subpopulations from the total population of LGR5+ cells).
To test whether differences in cell cycle [a known factor influencing the sensitivity of cells to DNA damage (Ren et al, 2002)] would correlate with increased DNA damage sensitivity of LGR5hi cells compared to LGR5lo cells, different cell cycle markers were employed. Staining for Ki‐67 and PCNA did not reveal significant differences in cell cycle activity of position 1 and 2 cells compared to position 4 cells (Supplementary Fig S5A–D). Similarly, FACS analysis of freshly isolated LGR5+ ISPCs from non‐irradiated and irradiated (24 h after IR) LGR5‐GFPki mice did not reveal significant differences in the cell cycle profile of LGR5hi cells compared to LGR5lo cells (Supplementary Fig S5E–I). Together, these data indicated that cell cycle differences did not account for the increased DNA damage sensitivity of LGR5hi cells compared to LGR5lo cells.
Enhancement of p53‐dependent apoptosis in response to irradiation in LGR5hi cells compared to LGR5lo cells
LGR5hi cells and LGR5lo cells from 3‐month‐old mice were FACS‐purified 6 h after IR. Gene expression array analysis of non‐irradiated and 12 Gy γ‐irradiated mice (6 h after IR, n = 4–5 samples per group) showed a significant increase in the number of genes that were induced or suppressed in response to γ‐irradiation in LGR5hi cells compared to LGR5lo cells (2,294 versus 567, P = 1.3 × 10−25, Fig 6A). A total of 266 genes were regulated in both cell populations in response to IR. However, the level of IR‐induced changes in the expression of these genes was significantly higher in LGR5hi cells compared to LGR5lo cells (median absolute fold change 4.9 versus 3.8, P < 0.0001, Fig 6A). Among the genes that were more strongly induced in LGR5hi cells versus LGR5lo cells was a set of apoptosis‐inducing genes including the p53 targets PUMA, NOXA, and BOK. qPCR analysis confirmed that IR led to a stronger induction of these pro‐apoptotic genes in LGR5hi cells compared to LGR5lo cells (Fig 6B). Enhanced activation of p53‐dependent apoptosis could explain the increased sensitivity of LGR5hi cells to DNA damage compared to LGR5lo cells. In agreement with this interpretation, Western blot analysis of freshly isolated LGR5+ cells derived from mice exposed to 12 Gy γ‐irradiation (6 h after IR) revealed an enhanced expression of phosphorylated p53 and cleaved caspase‐3 in LGR5hi cells compared to LGR5lo cells (Fig 6C, Supplementary Fig S6). TUNEL staining for apoptotic cells at 3 h after IR depicted reduced rates of apoptosis in position 4 cells compared to position 1 and 2 cells (Fig 6D and E). To test the functional role of p53 in enhancing radio‐sensitivity of LGR5hi cells compared to LGR5lo cells, the depletion of ISPCs was determined in p53−/− compared to p53+/+ LGR5‐reporter mice. p53 deletion diminished IR‐induced depletion of the total number of LGR5+ cells (Fig 6F), but the rescue of IR‐induced depletion of LGR5hi cells was more pronounced compared to the rescue of IR‐induced depletion of LGR5lo cells (Fig 6G, I, K, M and O). Gating of sub‐populations within the population of LGR5hi cells showed that the rescue in radio‐sensitivity in response to p53 deletion was much more pronounced in LGR5hi‐high cells compared to the LGR5hi‐low cells (Fig 6H, J, L, N and P, see Fig 1F for gating of subpopulations from the total population of LGR5+ cells). Together, these results indicated that p53‐dependent apoptosis contributes to the enhanced radio‐sensitivity of ISPCs with high LGR5 expression compared to ISPCs with low LGR5 expression.
Instructed modification of canonical Wnt/β‐catenin signaling activity changes the radio‐sensitivity of ISPCs in culture
The experiments on irradiated mice had shown a transient increase in the total number of LGR5+ cells in intestinal basal crypts at early time points after IR (3 h after IR), suggesting that Wnt signaling was transiently upregulated at early time points after IR (Fig 5O). To confirm these results, Wnt signaling activity was analyzed in ISPCs of cultured crypts derived from 3‐month‐old LGR5‐GFPki mice at 3 and 12 h after IR. Both LGR5hi and LGR5lo cells exhibited a significant induction of Axin2 at 3 h after IR compared to non‐irradiated controls, but the level went back down at 12 h after IR (Supplementary Fig S7A). Together with the data on enhanced p53 activation in LGR5hi cells compared to LGR5lo cells (Fig 6A–C), the data on transient upregulation of Wnt signaling in response to IR suggested that DNA damage induces an activating feed‐forward loop involving a transient upregulation of Wnt signaling, which in turn amplifies DNA damage responses, thus sensitizing ISPCs with intrinsically high Wnt activity to undergo DNA damage‐induced depletion. According to this model, an activation or inhibition of Wnt signaling should lead to respective changes in the sensitivity of ISPCs exposed to DNA damage. To test this assumption, freshly isolated crypts were cultured and transiently exposed to modifiers of canonical Wnt signaling shortly before IR. To inhibit Wnt signaling, recombinant DKK1 protein was added to the culture medium or the concentration of R‐spondin in the culture medium was reduced by 50% compared to normal conditions (Supplementary Fig S7B). To activate Wnt signaling, the GSK3 inhibitor 6‐BIO was added to the culture medium (Supplementary Fig S7C). Neither the activation nor the inhibition of Wnt signaling resulted in significant changes in cell cycle activity of LGR5+ cells in the crypt cultures (Supplementary Fig S7D and E). Of note, inhibition of Wnt signaling resulted in a significant decrease in radio‐sensitivity of both LGR5hi cells and LGR5lo cells (Fig 7A–F and K–M). Reversely, activation of Wnt signaling by treatment with 6‐BIO led to a significant increase in radio‐sensitivity of both LGR5hi cells and LGR5lo cells compared to control‐treated cultures (Me‐BIO) (Fig 7G–J and N–P).
Modification of canonical Wnt/β‐catenin signaling activity changes the radio‐sensitivity of ISPCs in vivo
To test whether the modulation of Wnt signal activity would also change the radio‐sensitivity of ISPCs in vivo, anti‐LRP6 antibody was injected to mice shortly before irradiation to transiently inhibit Wnt signaling (Ettenberg et al, 2010). Anti‐LRP6 antibody injection led to a significant reduction in GFP expression in crypt base stem and progenitor cells (Fig 8A–G) and in Axin2 expression in freshly isolated Lgr5hi cells compared to vehicle control‐injected mice (Supplementary Fig S7F). Notably, single dose of anti‐LRP6‐antibody injection did not lead to a significant loss of bottom ISPCs though lowering the GFP expression level in these cells (Fig 8H, I and L). Inhibition of Wnt signaling rescued the maintenance in the total number of crypt bottom PCNA+ ISPCs in the intestinal epithelium of irradiated mice but most pronounced in the survival of cells in position 1 and 2 of the basal crypt (Fig 8J–L). FACS analysis showed that anti‐LRP6 antibody treatment led to a reduction of LGR5hi cells and an increase of LGR5lo cells in non‐irradiated mice resulting in an improved maintenance of LGR5+ cells after irradiation (Fig 8M–R).
To analyze consequences of Wnt activation on IR‐induced depletion of ISPCs, APCmin/+ mice with enhanced Wnt activity (Zeilstra et al, 2008) were crossed to LGR5‐GFP reporter mice. APCmin/+ mice exhibited an enhanced GFP expression intensity in position 1–4 cells and even some GFP expression in TA cells above position 4 (Fig 9A–C). Notably, activation of Wnt signaling by genetic mutation of APC led to a greater loss of position 1–4 ISPCs upon irradiation (Fig 9D–F). FACS analysis also showed that IR led to a significantly enhanced depletion of LGR5+ cells in the intestinal epithelium of APCmin/+ mice compared to APC+/+ mice (Fig 9G–K).
The current study provides the first experimental evidence that cell intrinsic levels of canonical Wnt signaling determine heterogeneity in the sensitivity of ISPCs to DNA damage. These findings could be relevant for the evolution of stem and progenitor cell dysfunction and the selective survival of aberrant and mutant stem and progenitor cell clones in conditions associated with DNA damage accumulation such as aging and carcinogenesis.
Wnt signaling and stem and progenitor cell aging
This study shows that DNA damage results in the depletion of stem and progenitor cells with intrinsically high levels of Wnt signaling in the intestinal epithelium. There is increasing evidence that Wnt signaling, in addition to being an important factor for self‐renewal of stem cells and tissue regeneration, can also contribute to tissue aging. Along these lines, it was shown that over‐activation of Wnt in transgenic mice leads to defects in HSC functionality (Kirstetter et al, 2006; Scheller et al, 2006) and exerts deleterious effects on the maintenance of epithelial stem cells (Castilho et al, 2009). Systemic increases in Wnt signaling led to premature aging in Klotho‐deficient mice (Liu et al, 2007), and parabiosis studies revealed evidence that aging‐associated increases in Wnt stimulatory activity in blood serum contribute to the decrease in stem cell functionality and impaired regeneration during aging of wild‐type mice (Brack et al, 2007).
Despite these evidences that increases in Wnt signaling have adverse effects on stem cell maintenance and tissue aging, the mechanism is still poorly understood. The current study indicates that DNA damage represents a major confounding factor limiting stem cell maintenance in the context of elevated Wnt signaling. The study shows that cell intrinsic heterogeneity in the level of Wnt activity in stem and progenitor cells modulates their sensitivity to DNA damage. High levels of Wnt signaling amplify p53‐dependent DNA damage responses and increase stem and progenitor cell apoptosis in response to DNA damage. These findings stand in agreement with previous studies showing that Wnt activity can amplify DNA damage responses in other cell types (Damalas et al, 1999; Xu et al, 2008; Zhang et al, 2011). The current study shows that DNA damage leads to an amplification of Wnt signaling in ISPCs at early time points after DNA damage suggesting a feed‐forward loop of DNA damage inducing Wnt signaling, which in turn amplifies the DNA damage response (DDR). This signal‐enhancing forward loop contributes to the induction of DNA damage checkpoints in ISPCs. In line with this interpretation, this study shows that elevating Wnt signaling increases radio‐sensitivity of ISPCs, while inhibition of Wnt signaling reduces the radio‐sensitivity of ISPCs both in vitro and in vivo.
This study supports a model indicating that stem and progenitor cells with intrinsically low Wnt signaling activity represent a backup population ensuring recovery of tissue maintenance and survival in response to acute or chronic DNA damage. It was demonstrated by live cell imaging studies that +4 cells consist of a mixture of early TA cells (LGR5‐GFP‐negative) and LGR5‐GFP+ stem cells (Ritsma et al, 2014). In addition, lineage‐tracing experiments revealed that Dll1+ TA cells revert to stem cells in response to severe tissue damage (van Es et al, 2012). However, LGR5+ cells were shown to be essential for survival of mice in response to IR, and reversion of Dll1+ TA cells into stem cells is not sufficient for mouse survival in response to IR (van Es et al, 2012; Metcalfe et al, 2014). The current study shows that within the fraction of LGR5+ cells, the cells with low LGR5 expression and low Wnt signaling activity (low Msi1, low Axin2) preferentially survive in response to DNA damage. This holds true for FACS‐gated subpopulation of LGR5‐high positive cells (LGR5hi‐high cells being more sensitive to IR than LGR5hi‐low cells) and coincides with preferential survival of GFP‐positive cells in position 4 of the basal crypt. Together, these results stand in accordance with the concept that niche‐dependent local signals and cell intrinsic Wnt signaling modulate the survival of ISPCs in response to DNA damage. Based on our study, it is conceivable that Wnt signaling has a dual role in maintenance of the intestinal epithelium in response to acute DNA damage. While the lowering of Wnt signaling can enhance the survival of ISPCs in response to DNA damage, the elevation of Wnt signals appears to be beneficial for the regenerative phase following the acute damage (Zhou et al, 2013). This latter process likely includes pro‐regenerative effects at the progenitor cell level. Along these lines, it was shown that Wnt activation promotes radio‐resistance of progenitor cells in primary mammary epithelial cell cultures (Woodward et al, 2007).
Wnt signaling and maintenance of stem cells and cancer stem cells
An activation of Wnt signaling represents one of the hallmark features of cancer (Polakis, 2000; Holland et al, 2013) including the development of malignancies in the hematopoietic system and the intestinal epithelium (Reya & Clevers, 2005). Studies on the role of Wnt signaling in the maintenance of adult, non‐transformed stem cells revealed that Wnt activity is required for stem cell maintenance, but over‐activation of Wnt leads to stem cell depletion (Reya et al, 2003; Fevr et al, 2007; Fleming et al, 2008; Luis et al, 2009; de Lau et al, 2011). These previous data point to the existence of checkpoints that limit self‐renewal of stem cells with abnormally increased Wnt activity. The current study shows that higher levels of Wnt signaling increase the sensitivity of stem and progenitor cells to DNA damage in the intestinal epithelium. Since DNA damage accumulates in response to proliferation of cancer stem cells (Viale et al, 2009), it is conceivable that Wnt‐induced amplification of DNA damage signals in cancer stem cells may act as a tumor suppressor mechanism. The over‐activation of Wnt signaling in cancers indicates that tumor stem cells escape Wnt‐dependent amplification of DNA damage signals possibly involving the depletion of DNA damage checkpoint genes, such as p53. Along these lines, it has been observed that p53 inactivation promotes the progression of tumors induced by enhanced Wnt signaling in mouse models and human cancer cells (Donehower et al, 1995; Halberg et al, 2000; Kim et al, 2011). Interestingly, interspecies comparison on mouse and human intestinal carcinogenesis supports a model indicating that an optimal level of moderately increased Wnt signaling promotes the formation of tumorigenic lesion while excessive high levels of Wnt activity do not (Leedham et al, 2013). It is tempting to speculate that Wnt‐dependent sensitizing of intestinal tumor stem cells to DNA damage could contribute to the selection of only moderately increased Wnt activity during tumor formation.
Current experiments support the notion that heterogeneity in Wnt signaling activity in non‐transformed ISPCs originates from the position of the stem cells in the stem cell niche. CBCs that are located in between the Paneth cells at the crypt base exhibit higher Wnt signaling than those cells located above the upper most Paneth cells flanked by only one or two Paneth cells. The increased DNA damage resistance of ISPCs in stem cell niches with reduced Wnt stimulatory activity could enable the survival of initiated tumor cells, which often exhibit an increase in DNA damage and an activation of DNA damage checkpoints (Ye et al, 2007). In addition, there is evidence that the tumor stroma influences heterogeneity in Wnt signaling activity in cancer cells (Vermeulen et al, 2010). It is tempting to speculate that the heterogeneity of Wnt activity in tumor cells would also influence the sensitivity of established tumors to DNA‐damaging agents.
Together, the current study provides the first experimental evidence that the level of Wnt signaling activity represents a major confounding factor modulating the sensitivity of ISPCs to DNA damage. Selective survival of stem and progenitor cells with low Wnt signaling activity could influence the selection of aberrant stem and progenitor cells in the context of DNA damage during aging and carcinogenesis. This study also shows that treatment‐induced modification of canonical Wnt signaling activity changes the radio‐sensitivity of ISPCs. These data support the concept that modulation of canonical Wnt signaling could represent a novel therapeutic target to influence the survival of non‐transformed stem cells and possibly also of tumor stem cells in response to DNA‐damaging agents.
Materials and Methods
Mice were bred and maintained in the animal facilities of Ulm University and Leibniz Institute for Age Research on a standard diet and on a 12‐h light–dark cycle. Mice were monitored by weekly inspection. All mice were maintained on a C57BL/6J background. mTerc−/− mice were intercrossed to generate G3 mTerc−/− mice. Lgr5‐EGFP‐IRES‐CreERT2 mice were kindly provided by Hans Clevers. Lgr5‐EGFP‐IRES‐CreERT2 mice were crossed with mTerc−/− mice and intercrossed to generate LGR5ki; G3 mTerc−/− mice. P53 knockout mice and APCmin/+ mice were bought from Jackson Laboratory, and they were crossed with Lgr5‐EGFP‐IRES‐CreERT2 mice, respectively. C57BL/6J wild‐type mice were from Janvier Labs, France. All mice experiments were approved by the state government of Baden‐Württemberg (Protocol Number 35/9185.81‐3/919) and Thüringen (Protocol Number 03‐006/13).
RNA isolation and cDNA synthesis
Total RNA was isolated from freshly sorted cells or crypts by using MagMAX™‐96 Viral RNA Isolation kit (Ambion). DNA digestion step was included during the RNA purification process. Reverse transcriptions were performed to synthesize first‐strand DNA by using the GoScript reverse transcription system (Promega).
Quantitative real‐time PCR
qPCR was performed with an ABI 7300 Real‐Time PCR System (Applied Biosystems) in duplicates or triplicates from at least three biological samples. Primer sets for the detection of single genes are listed in Supplementary Table S1. iTaq SYBR Green supermix with Rox (Bio‐Rad) was used. Expression of genes was normalized to GAPDH in each sample.
For LGR5‐GFPki mice crypt cell flow cytometry, crypt single‐cell suspension was prepared as described (Sato et al, 2009). For cell cycle analysis, Cytofix/Cytoperm Kit (BD Biosciences) was used. Data acquisition and cell sorting were performed on FACS LSRII and FACS AriaIII (BD Biosciences). Data were analyzed with the BD FACS Diva software (BD Biosciences).
5‐μm paraffin sections of small intestine were used for immunofluorescence staining. Antibodies used were the following: PCNA (Calbiochem), Ki‐67 (Monosan), GFP (SantaCruz), Msi1 (eBioscience), anti‐mouse‐IgG‐Cy3 secondary antibody (Invitrogen), anti‐mouse‐IgG‐FITC secondary antibody (Invitrogen), anti‐rat‐IgG‐FITC (Invitrogen), and anti‐rabbit‐IgG‐Cy3 secondary antibody (Invitrogen).
Restrict counting regime of ISPCs was performed as following: Positions were counted starting from the lowest position of the basal crypt, which was defined as position 1. The neighboring position above the last Paneth cell was defined as position 4. ISPCs of both sides of the crypt were counted. Staining intensity of GFP and Msi1 was measured by ImageJ software in the Cy3 channel and FITC channel, respectively.
In situ hybridization
Eight‐micrometer paraffin sections of small intestine were used for in situ hybridization. The staining was done as described (Sperka et al, 2011).
TUNEL assay was performed using the In Situ Cell Death Detection Kit, Fluorescein (Roche) according to the manufacturer's instructions.
Equal numbers of cells were FACS‐sorted and lysed with Laemmli sample buffer (50 mM Trisbase pH 6.8, 4% SDS, 0.04% bromophenolblue, 20% glycerol). Whole‐cell lysates were boiled and loaded on 10 or 12% SDS–PAGE gel for electrophoresis. Following antibodies were used for immunoblotting: phospho‐p53 (Ser15) (1:1,000; Cell Signaling), cleaved caspase‐3 (1:1,000; Cell Signaling), and beta‐actin (1:10,000; Sigma).
Crypt culture was performed as previously described (Sperka et al, 2011). Briefly, isolated crypts were resuspended in cold Matrigel (BD) containing Y27632 (Abcam), R‐spondin, mEGF (PeproTech), Noggin (PeproTech), and penicillin/streptomycin and plated in 24‐well plate at density of 200 crypts, 50 μl per well. The plate was incubated at 37°C for 5 min, and then, 500 μl of advanced DMEM/F 12 medium (Invitrogen) containing B27 (Invitrogen), N2 supplement (Invitrogen), and 1.25 mM N‐acetylcystein was added to each well to cover the Matrigel. In the culture system, final concentrations of the following components were R‐spondin 1 μg/ml, mEGF 50 ng/ml, and Noggin 100 ng/ml. Y27632 10 μM was included in the first 3 days after seeding. The medium containing growth factors was changed every 3 days, and crypts were passaged every 9 days. Modification of Wnt activity was performed on day 9, 16 h before irradiation. DKK1 protein (R&D Systems) was used at a final concentration of 500 ng/ml. Rspo was prepared as previously described (Sperka et al, 2011). Six‐BIO (Calbiochem) or Me‐BIO (Calbiochem) was dissolved in DMSO and used at a final concentration of 20 nM. Single cell culture was performed as previously described (Sato et al, 2011). Briefly, sorted single LGR5hi or LGR5lo cells were pelleted and embedded in Matrigel, followed by seeding on a 96‐well plate (round bottom). Culture medium as described above with addition of 100 ng/ml Wnt3a (R&D Systems) was overlaid. Growth factors were added every other day, and culture medium was changed every 4 days.
RNA quality was controlled by the 2100 Bioanalyzer (Agilent Technologies). Quality‐proved RNA with a RIN value higher than 9.0 was used for gene expression analysis, which was performed using the SurePrint G3 Mouse GE 8x60K (Design ID 028005) Microarray Kit (Agilent Technologies). 50 ng of each sample was labeled with the Low Input Quick Amp Labeling Kit (Agilent Technologies) according to the manufacturer's instructions. Slides were scanned using a microarray scanner (Agilent Technologies). All expression data were deposited in Gene Expression Omnibus (GEO accession number GSE49461).
The LRP6 neutralization antibody was kindly provided by Dr. Seth A. Ettenberga and Michael P. Daley and applied to mice according to previous publications (Ettenberg et al, 2010). Briefly, anti‐LRP6 antibody (BpAb A7/B2) was injected intravenously with a single dose of 5 mg/kg 12 h before irradiation. Equal amount of IgG antibody was injected to control mice.
SPSS 11.5 and GraphPad Prism 6 softwares were used for statistical analysis. The unpaired two‐tailed Student's t‐test was used to calculate P‐values for all datasets except for the microarray analysis where Wilcoxon test was used.
ST and DT performed and analyzed majority of all experiments, including crypt isolation and culture, LGR5hi and LGR5lo cell purification, RNA isolation and qPCR, flow cytometry, intestinal tissue staining and counting, irradiation, and Western blot. YM did anti‐LRP6 antibody injection and helped in FACS experiments. TS and OO gave suggestions and helped in some crypt isolation experiments. AL performed microarray experiment. VS provided technical assistance for mouse experiments. JK and HAK performed microarray analysis. MK and KLR conceived and designed the experiments. The manuscript was written by ST, MK, and KLR and commented on by all other authors.
Conflict of interest
The authors declare that they have no conflict of interest.
This work was supported by the German Federal Ministry of Education and Research BMBF within its joint research project SyStaR (KLR, MK, HAK), by the EU (ERC to KLR: 323136—Stem Cell Geronto Genes) and by the International Graduate School in Molecular Medicine Ulm, funded by the Excellence Initiative of the German Federal and State Governments (GSC270).
FundingGerman Federal Ministry of Education and Research BMBF
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