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Pancreatic plasticity: epigenetic mechanisms and connections to neoplasia

David, Charles J. PhD

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doi: 10.1097/JP9.0000000000000036
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Abstract

Introduction

In multicellular organisms, a single genome must specify dozens to hundreds of different cell types. This is accomplished by the selective activation of cell type-specific patterns of gene expression. Protein coding sequences comprise a very small fraction of mammalian genomes, approximately 1.5% in humans.[1] Embedded in the large swaths of intervening sequence is a wealth of information that specifies cell type specific gene expression.[2] These intergenic information repositories are known as enhancers,[2,3] which can regulate the expression of their target genes at megabase distances, and are in some instances separated from target genes by several other genes.[4] The distance over which enhancers can act appears to be limited by the existence of topological-associated domains, which are megabase-sized chromatin regions that are insulated from surrounding chromatin.[5] Under normal conditions, enhancers are unable to affect gene expression of genes lying outside of such domains.[6–8]

DNA-binding transcription factors (TFs) are the primary “readers” of this information. Cell identity and function are specified by the combined actions of TFs, which can be crudely classified as lineage-determining TFs (LDTFs) and signal-dependent TFs (SDTFs—see Ref. [4] for a more complete classification).[9,10] LDTFs typically show a very restricted pattern of expression, while SDTFs are more ubiquitous, but only active in the presence of a specific signal. TFs bind cooperatively to DNA to promote cell-type and cell-state specific gene expression. Critically, SDTFs and LDTFs cooperate to activate gene expression,[11] meaning that the signals to which a cell is exposed can alter the function of LDTFs (e.g., from promoting self-renewal to promoting differentiation[12]), while, the collection of LDTFs expressed in a cell profoundly affect the gene expression output of a given signal[13,14] (Fig. 1).

Figure 1
Figure 1:
Critical interactions between SDTFs and LDTFs. (A) SDTFs bind to distinct genomic sites in different cell lineages, depending on the cellular repertoire of highly expressed LDTFs. SDTFs and LDTFs show a high degree of overlap in DNA binding. (B) Signaling determines the cellular function of LDTFs through SDTF–LDTF cooperation. In the case of liver progenitors, the active YAP-TEAD TF complex cooperate with LDTFs Foxa2 and Hnf4a to bind progenitor genes. In adult hepatocytes, absence of YAP-TEAD results in Foxa2/Hnf4a redistribution. LDTF = lineage-determining transcription factor, SDTF = signal-dependent transcription factor.

Once cell type-specific gene expression programs are established, barriers inhibit cell type interconversion. In the mid-1900s, Conrad Waddington presented the concept of “canalization” to describe the specification of different cell types, which remains resonant in the molecular era.[15] In his conceptualization, a marble is placed on the top of a hill, representing the totipotent state. As this marble rolls down the hill, it encounters various forks, representing alternate cell fates. As it continues down a given canal, it may encounter additional fate choices, until it finally reaches a minimum, achieving a stable fate. This concept was prescient, as it envisioned epigenetic barriers separating alternative cell fates, analogized in the form of the barrier separating 2 canals.

More recently, the molecular basis of the barrier between cell identities has become clear. The most conspicuous of these barriers revolves around the fact that eukaryotic DNA is packaged around histone proteins to form nucleosomes. Nucleosomes form a natural impediment to the DNA binding of most TFs, and provide an important basis for cooperation between TFs. While an individual TF may have difficulty competing with a histone octamer for binding to its cognate DNA sequence, when motifs for several TFs are present within the same nucleosome-bound DNA fragment, coexpression of these TFs allows cooperative eviction of nucleosomes resulting in chromatin opening, the first step in enhancer activation.[2,9] This form of cooperation often requires the activity of ATP-dependent histone remodeling complexes such as the SWI/SNF complex that, when recruited by TFs, help to evict nucleosomes and facilitate binding of cooperating factors.[16] Histones can also be posttranslationally modified in numerous ways that contribute to activation or repression of transcription.[17] TFs work in concert with factors that modify both the histone proteins and DNA itself to lock in cell identity, contributing to the barrier between cell fates (canals) envisioned by Waddington.

The barriers between most pairs of cell types are never breached without experimental intervention, the most famous example of which is the reprogramming of fibroblasts to pluripotent stem cells by a cocktail of 4 TFs.[18] That said, in some settings cell identity conversion can occur. A common form of cell identity change in human tissues is known as metaplasia, in which 1 cell type is replaced by another cell type in a given tissue. Metaplasia often results from injury, and can be enhanced by an accompanying inflammatory response.[19] In some tissues, including the pancreas, metaplasia is a major pre-cancerous state. Metaplasia can occur through direct conversion of 1 cell type to another, and usually occurs between cell types that share a common progenitor cell. The mechanisms underlying such cell fate switches are only beginning to be understood. Given the transcription-centric and chromatin-centric nature of cell identity specification, it is clear that deregulation of these facets of gene expression control must play a central role in breaking down the barrier between 2 cell identities. Below I will discuss transcriptional control of cell identity in the pancreas, and the ways in which injury and oncogenic insults disrupt and coopt these processes.

Epigenetic control of cell identity in the endocrine pancreas

In contrast to tissues such as the skin and intestine, the adult pancreas undergoes relatively little cellular turnover. Instead of a dedicated pool of stem cells, the healthy pancreas relies largely on self-duplication to replace aging cells and maintain pancreatic mass.[20,21] This model for tissue homeostasis is distinct from the hierarchical organization of many highly proliferative tissues such as the intestine[22] or skin,[23] in which self-renewing multipotent stem cells at the top of the hierarchy produce all of the cell types present in the tissue. In the hierarchical model, terminally differentiated cells are non-dividing, and devoid of regenerative capacity.

The existence of a hierarchical arrangement in the pancreas was cast into doubt when lineage tracing demonstrated that new insulin-producing β cells are not produced from a non-β-cell source.[24] In this experiment, β cells were genetically labeled at a specific time point, such that labeled β-cell progeny would also be marked, a technique known as lineage tracing.[25] If β cells are produced from non-β cells, the proportion of labeled cells should decrease over time, as the unlabeled progeny of stem cells replace the labeled cells. Instead, the authors showed that the percentage of labeled β cells remained constant over time, arguing for β-cell duplication as the primary means of endocrine cell homeostasis.

In addition to ruling out a stem cell source of β cells, the results of this experiment suggest that epigenetic barriers exist such that under normal conditions, little interconversion between β cells and other cell types occurs. Given the centrality of LDTFs in specifying cell identity, their regulation is central to epigenetic mechanisms that maintain cell identity. In β cells, the α-cell master TF Arx is repressed through several mechanisms, including Dnmt1-dependent repressive promoter methylation.[26] This promoter methylation is a key epigenetic barrier between the 2 states; β-cell directed deletion of Dnmt1 resulted in the ectopic expression of Arx and conversion of β cells into α cells.[26] The β-cell master TF Nkx2.2 contributes to the epigenetic locking in of β-cell fate through binding of the Arx locus and the recruitment of the transcriptional corepressor Grg3, which in turn recruits histone deacetylases (HDACs), resulting in Arx repression.[27] Because diabetes is caused by β-cell loss, there is considerable interest in the possibility of ameliorating the condition by converting other types of cells into β cells. As suggested by the lineage tracing experiments outlined above, this is something that does not occur under normal homeostasis. Similar to the mechanisms preventing β-to-α conversion, the reverse conversion is reciprocally prevented by the combined actions of Arx and Dnmt1.[28] Loss of these 2 factors in α cells results in reversion to a β-cell like fate. Recent breakthroughs in β-cell replenishment have employed γ-aminobutyric acid (GABA) and artemisinins, which function through the GABA receptor to affect α-to-β-cell conversion in the pancreas.[29,30] In the case of artemisinins, this effect occurred through nuclear-to-cytoplasmic translocation of Arx.[29]

In contrast to normal homeostasis, the epigenetic barriers between β-cell identity and other cell fates can be lowered in response to insult. By targeting the diphtheria toxin receptor to the β-cell compartment in mice, Thorel et al[31] were able to rapidly ablate 99% of β cells upon diphtheria toxin administration. When provided insulin, the mice survived and showed a marked replenishment of β cells over time. The authors showed by lineage tracing that replenished β cells were derived from the α-cell compartment. In addition to this mechanism of β-cell replacement, it was later shown that somatostatin-producing δ cells can repopulate the β-cell compartment in younger mice.[32] Forced conversion of α cells to β cells has been shown to occur upon overexpression of the TFs Mafa, Pax4, or Pdx1,[33,34] something that has now been harnessed in pre-clinical gene therapy trials.[35]

What is the epigenetic basis for the plasticity that allows non-β endocrine cells to form β cells when they are depleted? Cooccurrence of H3K4me3, a mark associated with active promoters and repressive H3K27me3 renders target genes “bivalent.” Bivalent genes are not expressed, but are instead thought to be poised for activation in response to an appropriate signal. This phenomenon was first discovered in embryonic stem cells, where key lineage differentiation genes are silenced, but marked by bivalent histone marks, allowing them to be activated rapidly in response to signaling cues that promote lineage specification.[36] Examination of the distribution of H3K4me3 and H3K27me3 in purified endocrine cell populations has shed light on the plasticity of α cells, which were shown to contain a higher proportion of bivalent genes than β cells.[37] Importantly, many β-cell genes, including key LDTFs Mafa and Pdx1 were bivalently marked in α cells, suggesting that these genes are primed for activation in response to an appropriate signal. Consistent with the proposed role for H3K27me3 in gene repression,[38] the authors went on to show that inhibition of histone methyltransferase(s) responsible for the mark through the general histone methyltransferase inhibitor Adox ectopically activated β-cell genes within the α cells. Shared promoter DNA hypomethylation patterns between α cells and β cells, specifically at key, highly expressed, cell type specific genes such as insulin and glugacon also appears to play a role in plasticity between the 2 cell types.[39] In this case, the authors proposed that hypomethylation of β-cell specific genes in α cells licenses their expression in the event of a fate conversion, and is possibly a pre-requisite for plasticity. While the precise mechanism of the promoter demethylation was not identified, demethylation was observed to occur upon differentiation of Ngn3-positive endocrine progenitor cells, a precursor of both α cells and β cells. Demethylation is presumably driven by TFs expressed during this developmental window. These results shed light on mechanisms allowing plasticity cell types that share close developmental origins.

Endocrine neoplasia through epigenetic misregulation

Pancreatic neuroendocrine tumors (PanNETs) are the second most common form of pancreatic neoplasia and are frequently associated with hormone secretion (insulin, glucagon, gastrin), reflecting a perturbation of normal endocrine cell programs.[40] LDTFs that determine normal endocrine cell types were highly expressed in PanNETs in a manner corresponding to the secreted hormone,[41] further illustrating the likelihood that these tumors are an outgrowth of fully or partially differentiated cells of origin, a supposition that has been borne out in mouse models (discussed below). Hormone-secreting PanNETs tend to be less deadly than the more poorly differentiated neoplasms known as neuroendocrine carcinomas (NECs). NECs are more proliferative, and fail to produce hormones, leading to speculation that these tumors arise from a more primitive endocrine precursor.[40] PanNETs and NECs display a distinct mutational profile, indicating that they may indeed be distinct diseases.

Well differentiated, hormone-secreting PanNETs are frequently associated with the lineage-specific loss of several chromatin regulatory factors, including MEN1, ATRX, and DAXX.[42,43] The connection between MEN1 and predisposition to endocrine tumors has long been appreciated, as its mutation is connected with the familial syndrome that provided its name (multiple endocrine neoplasia).[44,45] The gene is mutated in a dominant inherited disorder that results in frequent endocrine adenoma formation in the pancreas and other endocrine organs.

Mouse models in which Men1 deletion is targeted to specific endocrine cell compartments have been developed to recapitulate PanNET formation. Interestingly, these models have provided additional examples of plasticity between α and β-cell fates, this time in a neoplastic context. Using a transgenic mouse in which Cre expression was targeted to α cells using the glucagon promoter, glucagonomas, and insulinomas were both frequently observed, with insulinomas coming to predominate over time.[46] Interestingly, coexpression of glucagon and insulin was observed in young mice, before tumor formation but concomitant with increased proliferation. The transdifferentiation did not always coincide with expression of a full complement of β-cell LDTFs, indicating that the transition may be to an aberrant β-cell-like entity rather than activation of a normal differentiation program.[46] Interestingly, later work indicated that the reverse transition is also possible when Men1 is deleted in β cells, which resulted mostly in insulinomas but also an appreciable rate of glucagonomas.[47] It is noteworthy that in human PanNETs, MEN1 alteration is more closely associated with an α-cell gene expression signature.[48] It is currently unclear how Men1 loss licenses lineage infidelity in the pancreatic endocrine compartment, and whether transdifferentiation, as opposed to distinct cells of origin, may explain the phenotypic differences in human functional PanNETs.

MEN1 lacks clearly discernable domains that might provide insights into its function, initially making its precise function within the cell challenging to identify. One route to identification of its function has been through identification of interacting proteins.[49,50] MEN1 is a predominantly nuclear protein, and its classification as a chromatin-associated modifier of gene expression was suggested by its interaction with numerous TFs and epigenetic regulators.[50] Notably, it was found to exist in a complex with trithorax family histone methyltransferases MLL2 and ASH2L, indicating a role in developmentally regulated gene expression.[51] Like many transcriptional regulators, the function of MEN1 in tumorigenesis is highly context-specific; despite its tumor suppressive activities in endocrine cells, MEN1 is essential for MLL-driven leukemia progression through an interaction with MLL,[52] and also promotes castration-resistant prostate cancer.[53] MEN1 has been proposed to function as a molecular scaffold, bridging interactions between signaling molecules and transcriptional regulators to affect transcription.[50] In contrast to its role in leukemia, in islets, the interaction of MEN1 with MLL, an H3K4 methyltransferase associated with gene activation, promotes recruitment of MLL to and activation of cyclin-dependent kinase inhibitors p27 and p18 to repress proliferation.[54] MEN1 interacts with JUND, repressing JUND-mediated transcriptional activation in a manner disrupted by disease-causing MEN1 mutations,[55] likely in part due to blockade of JUN N-terminal kinase-mediated JUND phosphorylation[56] and recruitment of the repressive mSin3a HDAC complex.[57] In addition, MEN1 interacts with a bevy of transcriptional effectors of various signaling pathways connected to cancer, including Smads, NFκB, and Wnt.[58] MEN1 interacts with β-catenin, a transcriptional effector of the Wnt pathway, and its loss was shown to promote aberrant β-catenin nuclear accumulation in a mouse model of β-cell-specific Men1 ablation.[59] MEN1-deficient PanNETs are dependent on Wnt/β-catenin signaling,[60] so this function of MEN1 appears likely to contribute to its context-specific tumor suppression. MEN1 also regulates the hedgehog (Hh) pathway by promoting inhibitory PRMT5-mediated histone arginine methylation at the promoter of GAS1, an Hh coreceptor, dampening the output of the pathway.[61] Inhibition of the Hh component smoothened (SMO) reduced proliferation in a mouse insulinoma model.[61]

In 2012, an exome sequencing study revealed ATRX and DAXX as 2 other frequently altered chromatin-associated proteins in PanNETs.[42] Unlike chromatin-associated proteins given the “epigenetic” label, ATRX and DAXX play important roles beyond the control of gene expression. ATRX and DAXX form a complex and specifically deposit histone H3.3 at repetitive DNA, notably at telomeres.[62,63] While the precise function of H3.3 at telomeres is still an open question,[64] mutational loss of ATRX and DAXX is strongly associated with abnormally long telomeres through a mechanism known as alternative lengthening of telomeres.[65] Loss of DAXX and ATRX is associated with a higher degree of chromosomal instability in PanNETs, and reduced patient survival.[66] It remains to be seen whether the functions of ATRX and DAXX at telomeres is indeed the primary driver of tumorigenesis spurred on by their absence. It should be noted that the loss of Atrx in a different context, glioma-initiating cells of the mouse brain, leads to widespread changes in chromatin accessibility and gene expression, and leads to alterations in cell identity.[67] In light of these observations, it is likely too early to conclude which functions of these histone chaperones is most relevant to tumor suppression.

Pancreatitis, plasticity, and progenitor cells in the pancreas

Given the importance of β-cell depletion to human disease, there has been a strong interest in the possibility that stem or progenitor cells might exist as a source of new β cells, known as neogenesis. The existence of stem or progenitor cells in the pancreas has been a controversial topic, and the reader is referred to several recent reviews for a more in-depth discussion.[20,68,69] Ductal cells had been considered an attractive candidate pool of potential progenitor cells for β-cell neogenesis, an idea supported by an early lineage tracing study.[70] However, subsequent work has not supported a role for postnatal ductal cells in the formation of endocrine cells under normal circumstances.[20]

When the pancreas sustains various forms of injury, an inflammatory condition known as pancreatitis results.[71,72] In humans, pancreatitis is associated most strongly with alcoholism and smoking, in addition to a much more rare hereditary pancreatitis predisposition. Pancreatitis appears most likely to result from a blockage of normal pancreatic enzyme secretion that can occur through the obstruction of normal pancreatic ductal flow. This can result in the inappropriate release of activated digestive enzymes in the pancreatic interstitial space, resulting in injury to surrounding cells, causing an inflammatory response.[71] The major histopathological result of this sequence of events is atrophy of the acinar cell mass, and the appearance of tubular structures reminiscent of ductal cells. The role of inappropriately activated digestive enzymes in the etiology of the disease is underscored by the fact that hereditary pancreatitis can be caused by mutations in PRSS1, the gene encoding cationic trypsinogen.[72] The pancreatitis-associated mutations invariably lead to inappropriate autocatalytic activation of the zymogen or stabilization of the active enzyme.

In rodents, there are 2 major models that recapitulate aspects of human pancreatitis. In one, partial duct ligation (PDL), the major pancreatic duct that drains the tail of the pancreas into the duodenum is ligated, forming a blockage similar to gallstone pancreatitits.[73] This results in massive acinar apoptosis and inflammation.[74,75] A second model involves the administration of a choleocystokinin analog known as caerulein, most commonly through intraperitoneal injection. Caerulein is a secretagogue that stimulates the inappropriate release of proteolytic enzymes from acinar cells, thus mimicking the etiology of human disease.[76] As with PDL, caerulein administration leads to massive acinar apoptosis, accompanied by the appearance of ductal cells.

Mouse models of pancreatitis have shown conclusively that the exocrine pancreas is capable of a high degree of plasticity upon inflammatory insult.[20,77,78] Lineage tracing has demonstrated that ductal structures observed in the inflamed pancreas can result from transdifferentiation of preexisting acinar cells into a duct-like state expressing markers such as Krt19, a process known as acinar-to-ductal metaplasia (ADM).[77,79] It is now clear that these cells are of critical importance to both normal pancreatic regeneration and tumorigenesis.[77,80] While normal ductal cells are incapable of serving as endocrine progenitor cells, there is evidence that the transdifferentiated duct-like cells within ADM lesions are capable of serving as endocrine progenitors.[81] Clearly the duct-like state achieved in ADM is a progenitor-like state quite distinct from normal ducts.

Acinar-to-ductal plasticity as a prelude to cancer

Pancreatic ductal adenocarcinoma (PDA) is the most common form of pancreas cancer, and it consistently ranks as the tumor type with the worst prognosis.[82–84] The most common precursor lesions for PDA are pancreatic intraepithelial neoplasias (PanINs), which are graded PanIN-1, PanIN-2, and PanIN-3 depending on their degree of atypia.[85] Epidemiological studies have established a link between chronic pancreatitis and increased risk for PDA.[86] Congruent with this, studies in mice that have demonstrated that metaplastic ductal cells are easily transformed by mutant Kras, and serve as a major cell of origin for PanINs.[87,88] These findings are based on the observation that acinar cells, rather than ductal cells serve as a more efficient cell of origin for PanINs.[89,90] While these results might have been initially puzzling given the ductal histology of PDA, the requirement for ADM as an early step in acinar-derived pancreatic tumorigenesis seems to explain the apparent discrepancy, and emphasizes the differences in oncogenic potential between normal ductal cells and those comprising ADM lesions (Fig. 2).

Figure 2
Figure 2:
Plasticity and tumorigenesis in the pancreas. Pancreatitis leads to ADM, and the emergence of duct-like cells that are sensitive to mutant Kras-driven transformation. In the presence of mutant Kras, ductal phenotype is locked in and PanINs develop. In the wild-type pancreas, the normal pancreatic structure is rapidly regenerated after removal of injury stimuli. ADM = acinar-to-ductal metaplasia.

The role of Kras mutations in the process of producing acinar-derived PanIN lesions appears to be 2-fold. While mutant Kras expression throughout the pancreas in young mice leads to little histological abnormality, mutant Kras is able to cooperate with caerulein treatment to drive a more complete ADM than that observed in Kras wild-type animals.[88,91] In wild-type animals, caerulein-induced ADM resolves within 1–2 weeks of cessation of drug treatment, likely through the redifferentiation of the transient ductal structures into the acinar cells that comprise the bulk of the normal pancreas. In a stark manifestation of its oncogenic abilities, the presence of mutant Kras completely blocks this redifferentiation, preserving the ductal identity of ADM lesions, which eventually give rise to PanINs.[88] This process occurs spontaneously at a low rate in mice harboring pancreatic expression of mutant Kras, as the incidence and grade of PanINs increases over time without caerulein administration.[92]

Transcriptional determinants of cell identity in the exocrine pancreas

To fully understand the process by which 2 cell types interconvert, it is first necessary to understand the transcriptional underpinnings of each cell state. Key LDTFs that specify the acinar and ductal states have been identified. Acinar identity is specified in part by pancreas-specific TF 1a (Ptf1a), a basic helix-loop-helix (bHLH) TF that is expressed in early pancreatic buds at embryonic day 9,[93] becoming restricted to the exocrine pancreas at later embryonic stages, where it is required for specifying the exocrine fate.[94] Ptf1a is expressed in multipotent progenitor cells of the pancreas during embryogenesis, while Ptf1a-expressing cells only give rise to acinar cells in postnatal mice (with the exception of injury, as discussed above).[81] Ptf1a functions as part of a trimeric complex consisting of another bHLH and either Rbpj (at early stages of pancreatic differentiation), or Rbpjl, as cells commit to an acinar cell fate.[95] The Rbpjl-containing complex confers characteristics of terminally differentiated acinar cells, driving expression of digestive enzymes.[95] Another acinar bHLH LDTF is Mist1 (Bhlha15), is required for normal differentiation and function of acinar cells.[96,97] Mist1 appears to play an important role in organizing secretory programs in a variety of specialized secretory cells in addition to pancreatic acinar cells.[98] A third major transcriptional determinant of the acinar state is Nr5a2, which is also required for normal acinar differentiation and function.[99–101] As is usually the case for LDTFs, extensive cooperation cross-regulation occurs between these 3 acinar state-determining factors, as the 3 factors cooperate to activate shared acinar-specific target genes,[99,102,103] and reciprocally activate each other's expression.[102,103] This feedforward mechanism presumably stabilizes and reinforces the acinar cell fate.

Ample evidence points to acinar LDTFs as a key epigenetic barrier that restrains a latent tendency of acinar cells to undergo ductal transdifferentiation. An early demonstration of this came by disruption of Mist1 function through transgenic expression of a dominant negative form of the protein, which led to activation of ductal genes in acinar cells, reminiscent of ADM.[104] In the presence of stimli that promote ADM, the phenotype of Mist1 loss of function is further compounded. In Mist1 knockout mice, expression of mutant Kras is greatly more effective in inducing PanIN formation from acinar cells.[105] A similar phenomenon is observed when Ptf1a and Nr5a2 are depleted.[100,101,106,107] Based on these results it is clear that acinar cells are primed to adopt a ductal fate upon loss of key acinar factors, or through induction of an appropriate inflammatory signal, while combination of the 2 further exacerbates the phenotype. Recent work has further elucidated the molecular switch between the 2 fates. Pancreata of Nr5a2−/+ mice are highly sensitive to induction of ADM and oncogenic transformation,[107] and exist in a constitutive pre-inflammatory state. Examination of gene expression changes that underlie this pre-inflammatory state in Nr5a2−/+ pancreata revealed changes in a number of inflammatory genes and increased activity of activator protein-1 (AP-1) family TFs, a transcriptional state that could be phenocopied in wild-type mice by caerulein administration.[101] Importantly, Nr5a2 itself is proposed to be intimately involved in the switch between terminal differentiation and inflammation, as it was shown that in Nr5a2−/+ mice the remaining Nr5a2 protein was found not at the usual acinar differentiation genes, but at the very inflammatory genes that higher levels of the protein apparently repress. This redistribution, known as the Nr5a2 “transcriptional switch” appears to occur through interactions between Nr5a2 and AP-1 family TFs.[101] It remains to be shown whether Nr5a2 actually drives the expression of inflammatory genes during this switch, or is simply present at their genes as a bystander. Arguing for the latter, mice entirely deficient for pancreatic Nr5a2 are highly prone to ADM and inflammation, indicating that the inflammatory gene expression program is not dependent on Nr5a2.[100] In any event, it is clear that Nr5a2 and other acinar identity factors actively repress a latent inflammatory and ductal identity program that exists in acinar cells.

While the acinar transcriptional program is silenced during ADM, a duct-like program must simultaneously be activated. Consistent with this, a number of ductal TFs are upregulated concomitant with silencing of the acinar gene expression program, and have been shown to be essential for ADM. Sox9 is an essential component of the embryonic pancreatic progenitor transcriptional program.[108] Sox9 expression becomes limited to ducts in adults, and it is required for ductal differentiation.[109] Sox9 is upregulated during caerulein-induced ADM, and, while not strictly required for ADM, is essential for KrasG12D-mediated reprogramming of these lesions into PanINs.[89] Overexpression of Sox9 in acini results in the ectopic activation of ductal genes, and cooperates with inflammation to potentiate ADM.[89] A similar essential role in ADM has been established for a set of additional lineage-restricted TFs, including Klf4,[110] Klf5,[111] and Etv5.[112] Pdx1 is a key determinant of embryonic pancreatic progenitors, and its overexpression can promote ADM.[113] Paradoxically, Pdx1 deletion in adult acinar cells promotes increased ADM, indicating that Pdx1 also plays a role in maintaining acinar identity.[114] However, Pdx1 upregulation is regularly seen in PDA and its associated pre-neoplastic lesions, indicating it may play a distinct role upon establishment of ductal identity. Indeed, Pdx1 was shown to be required for proliferation of several mouse and human pancreatic cancer cell lines.[114] Such functional switches for important LDTFs within different cells in the same tissue have been shown in other tissues.[12] In addition to lineage-type factors, stress-responsive and signal-responsive TFs also play a role in the transition to a ductal phenotype. Pancreatic injury that results in pancreatitis is coupled to activation of the unfolded protein response,[115] which in turn activates Atf3, which plays a role in repressing the acinar cell transcriptional program during ADM.[116] Il6-gp130 signaling plays an important role in the development of pancreatic cancer,[117] and a key transcriptional mediator of this pathway is Stat3, which is tyrosine phosphorylated in response to Il6 signaling. Stat3 knockout attenuated ADM and PanIN formation, and was required for progression to PDA.[118] Stat3 physically associates with another SDTF that is activated during inflammation, Nfatc1 to promote KrasG12D-promoted tumorigenesis to activate transcription of the Sox9 gene, providing an important link between the induction of inflammation and the activation of ductal identity. Finally, as mentioned above, activation of AP-1 family TFs during inflammation is required to drive ADM.[101]

Although many of the transcriptional players involved in the ADM–PanIN progression have been identified (the cast), we are still lacking a global picture of how they interact to drive the process (the play). Many outstanding questions remain. For example, the assumption of ductal identity by acinar cells apparently gives rise to cells primed for mutant Kras-driven neoplastic transformation, and many components of the ductal transcriptional network are required for this. What distinguishes the transformable duct-like cells in ADM from normal duct cells that are resistant to Kras-driven transformation? How does mutant Kras signaling interact with transcriptional network in the transformable ADM lesions to “lock in” the ductal phenotype? While normal ducts are resistant to Kras-driven transformation, normal ducts can give rise to PDA when Kras activation is coupled to p53 loss or mutation,[119,120] or, as will be discussed below, upon loss of chromatin remodeling factors. Interestingly, in this setting, a direct progression to PDA apparently occurs without a PanIN intermediate.[120] How p53 governs the transformability of normal ductal cells is currently unknown.

Epigenome reprogramming in PDA identity and progression

Cancer is, for good reason, considered a disease of the genome, as its initiation and progression is almost invariably caused by mutational activation of oncogenes and genetic loss of tumor suppressors. However, in recent years, limits to the role of genetics in determining tumor phenotype have become clear. Most notably, despite numerous attempts to identify them, no recurrent metastasis-driving genetic aberrations have been found in human samples.[121] This has required a search for epigenetic explanations for the acquisition of metastatic traits. Recent work has explored this using mouse organoids derived from the Pdx1-Cre; lox-stop-lox KrasG12D; lox-stop-lox p53R172H (KPC) model. Organoids derived from non-metastatic primary tumors show little ability to colonize the lung or liver, and grow relatively slowly when orthotopically transplanted. In contrast, organoids derived from lung, liver, or peritoneal metastases perform efficiently in metastatic colonization assays and grow rapidly at the primary site, indicating a general, catastrophic gain in aggressiveness in the metastatic tumors.[122] To probe epigenomic differences between the more indolent primary tumors and metastases, Vakoc et al subjected the 2 sets of organoids to enhancer profiling using chromatin immunoprecipitation sequencing for H3K27ac and H3K4me1, the combined presence of which mark enhancers. This resulted in strikingly reproducible patterns of hundreds of gained and lost enhancers in metastatic organoids compared to primary tumor organoids.[123] The gained enhancers were associated with an embryonic endodermal progenitor transcriptional signature, suggesting a dedifferentiation event as a driver of gained aggressiveness. The authors focused on the transcriptional determinants of gained enhancers, and searched for TF binding motifs enriched in these sites. Using this approach, they identified a forkhead box family motif and GATA-type motif. Members of both families were upregulated in metastatic organoids, namely, Foxa1 and Gata5. Overexpression of these factors in indolent primary tumor organoids was sufficient to recapitulate the progenitor-like enhancer and gene expression signature, and greatly boosted their metastatic properties.[123] Foxa1 and Gata5 likely represent only part of the story, as other motifs, most notably the AP-1 family motif, were even more enriched in the gained regions. Gains in mutant Kras signaling output have been shown to drive metastasis in PDA mouse models,[124] indicating that increased signaling through a Kras-AP-1 axis might also contribute to the gained metastatic abilities of the cells.

Similar approaches have been taken to explore the transcriptional basis of intertumoral heterogeneity in human cells. Comparison of enhancer landscapes in cells derived from low-grade and high-grade PDA revealed a set of grade-specific enhancers.[125] Interestingly, a KLF5-dependent program dominated low-grade specific enhancers, and this was lost in high-grade enhancers. Low-grade cells were dependent on KLF5 for maximal proliferation, while expression of KLF5 was lost in higher grade tumors.[125] As discussed above, Klf5 is required for PDA initiation of tumorigenesis in the mouse model.[111] It is also a determinant of the epithelial state that is downregulated during epithelial-to-mesenchymal transition (EMT), resulting in apoptosis in bulk epithelial tumor cells.[126] These results suggest that, while early events in pancreatic tumorigenesis are dependent on KLF5, cells in advanced tumors are able to find viable transcriptional configurations that are independent of KLF5, allowing them to be freed of the constraints on invasiveness imposed by the KLF5-dependent epithelial phenotype. This theme has emerged in other epithelial tumor types in which LDTFs that play a role in proliferation in the normal progenitors are required for early lesions and well differentiated tumors, while their loss is more advanced tumors is associated with increased invasiveness. Examples of this are GATA3 in breast cancer[127] and NKX2-1 in lung.[128] It should be noted that the organoid-based study of Vakoc et al[123] found no evidence of a contribution of EMT to increased aggressiveness in KPC tumors.

Transcriptional determinants that distinguish pancreatic cancer subtypes from each other have only very recently begun to be explored. The squamous subtype of pancreatic cancer represents approximately 25% of PDA, and has a particularly dismal prognosis.[129] The squamous subtype is associated with mutations in the H3K27me3 demethylase KDM6A,[129] and inactivation of Kdm6a combines with mutant Kras to drive squamous-type carcinomas in the murine pancreas.[130] Examination of the enhancer landscape in squamous compared with non-squamous PDA cells revealed a set of over 1000 squamous-specific enhancers, a program controlled by the squamous LDTF ΔNp63.[131] ΔNp63 conferred squamous characteristics on non-squamous tumors, and became a tumor dependency in squamous tumor cells. These results show that adenocarcinoma cells are capable of squamous transdifferentiation, indicating that squamous subtype tumors may emerge from the same cell of origin and precursor lesions as other PDA subtypes, which appears likely because the normal pancreas contains no squamous epithelia. The exact molecular events that surround the emergence of ΔNp63 expression in the squamous subtype are still unclear.

Chromatin regulators in pancreatic plasticity and tumorigenesis

A wide variety of cancers display frequent genetic alterations in chromatin remodeling factors, most notably members of the SWI/SNF complex.[132–134] In pancreatic cancer, ARID1A is the most frequently mutated SWI/SNF subunit.[135,136] Recent work in mouse models has begun to elucidate the role of key SWI/SNF subunits in pancreatic tumorigenesis. In the first direct demonstration of the tumor suppressive role of SWI/SNF in PDA, pancreas-specific knockout of the ATPase subunit Brg1 (Smarca4) was shown to promote the formation of ductal cell-derived intraductal papillary mucinous neoplasia (IPMN) a cystic neoplasm that can, like PanINs, serve as a precursor lesion for PDA.[137] Like human IPMN-associated PDA, the PDA formed in Brg1 conditional knockout mice was more indolent than PanIN-associated PDA driven by p53 knockout. The relationship between Brg1 loss and tumorigenesis is context-specific, because while it restrains neoplastic transformation of ductal cells, its loss in acinar cells reduced PanIN formation.[137] When Brg1 was restored in Brg1-null PDA cells, its tumor-suppressive properties were no longer apparent, and instead the cells displayed a more aggressive EMT-like phenotype.[138] Loss of Arid1a partially phenocopies loss of Brg1, as Arid1a loss in ducts results in an IPMN–PDA sequence that mirrors the effect of Brg1 deletion.[139,140] Distinct from Brg1, loss of Arid1a in acinar cells resulted in increased ADM and PanIN formation.[139,141,142] SWI/SNF complexes are large, multiprotein assemblies, that display developmental stage-specific and tissue-specific subunit composition.[143] Many SWI/SNF subunits enter the complex in a mutually exclusive manner (e.g., Arid1a or Arid1b). It is possible that the apparently disparate roles of Brg1 and Arid1a in acinar-derived PanIN formation is the result of tumor suppressive functions for Arid1a-containing complexes, while complexes lacking Arid1a but containing Brg1 play essential role in the ADM–PanIN sequence. Much work is necessary to explore the potential distinct roles of various SWI/SNF complexes during tumor formation. Interestingly, using a tetracycline-dependent short hairpin RNA (shRNA) to deplete Arid1a in vivo, the ADM phenotype was shown to be highly dependent on the timing of Arid1a inactivation with respect to Kras mutation. When Arid1a was depleted simultaneous with Kras mutation (on embryonic day 7), little ADM and PanIN formation was observed. In contrast, when Arid1a was depleted postnatally PanINs formed efficiently.[142] In contrast to irreversible loxp-mediated gene inactivation, inducible shRNAs also allow for the restoration of tumor suppressors upon tumor formation.[144] Using this approach, it was shown that, in contrast to tumor suppressors for which restoration results in regression of established tumors (such as p53[145] and adenomatous polyposis coli [APC][146]), Arid1a restoration had no effect in PanINs promoted by its loss.[142] This would be consistent with Arid1a restraining a transition to a transcriptional state that, once achieved, cannot be reversed.

Relatively few insights into the mechanisms by which loss of SWI/SNF components drives the formation of IPMN from ducts have been obtained. IPMN formation is thought to involve a still poorly defined “dedifferentiation” of ductal identity. In this process, ductal markers are downregulated while “progenitor” markers are upregulated, a process that occurs in ductal cells upon knockout of either Brg1[138] or Arid1a.[140] At least in the case of Arid1a, it has been suggested that this occurs through downregulation of Sox9,[140] although we are far from a mechanistic understanding of the tumor suppressive role of chromatin remodelers in pancreatic tumorigenesis. Attempts to directly look at the impact of Arid1a depletion on transcriptional programs in the pancreas using Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq) in sorted epithelial cells have been hampered by the high degree of epithelial heterogeneity in the inflamed pancreas.[142] Because of this, ATAC-seq results might simply reflect the degree of ADM in the 2 conditions, but shed no light on the relevant changes in the incipient transformed cell. An understanding of the effect of Arid1a depletion would require direct examination of purified tumor-initiating cells, something that is currently technically difficult.

Future prospects

Considerable progress has been made in elucidating the transcriptional underpinnings of cellular plasticity and tumorigenesis in the pancreas. Key factors contribute to both processes, providing clear molecular links between processes whose functional connections have long been clear. Dynamic changes in the LDTFs that determine cell identity are central to these processes in both the exocrine and endocrine pancreas. While many TFs that drive key pathophysiological processes from ADM to tumor formation and progression have been identified, systematic approaches will be necessary to a complete picture of the transcriptional networks that drive these processes, and especially the interplay between SDTFs and LDTFs. This will demonstrate how oncogenic signals tap into transcriptional programs that exist in the inflamed pancreas. Because all transcriptional programs are strongly dependent on and influenced by chromatin regulators, the impact of key factors, such as Arid1a, will need to be understood through their impact on essential transcriptional programs at a global and biochemical level. New techniques allowing the interrogation of small numbers of cells derived from in vivo sources, such as ATAC-seq should soon allow for this.

Acknowledgments

Work in the David laboratory is supported by a start-up package from Tsinghua University and the Peking University-Tsinghua Center for Life Sciences.

Author contributions

None.

Financial support

None.

Conflicts of interest

The authors declare no conflicts of interest.

References

1. Lander ES, Linton LM, Birren B, et al. Initial sequencing and analysis of the human genome. Nature 2001; 409:860–921.
2. Long HK, Prescott SL, Wysocka J. Ever-changing landscapes: transcriptional enhancers in development and evolution. Cell 2016; 167:1170–1187.
3. Rickels R, Shilatifard A. Enhancer logic and mechanics in development and disease. Trends Cell Biol 2018; 28:608–630.
4. Lettice LA, Heaney SJ, Purdie LA, et al. A long-range Shh enhancer regulates expression in the developing limb and fin and is associated with preaxial polydactyly. Hum Mol Genet 2003; 12:1725–1735.
5. Dixon JR, Selvaraj S, Yue F, et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 2012; 485:376–380.
6. Dowen JM, Fan ZP, Hnisz D, et al. Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes. Cell 2014; 159:374–387.
7. Flavahan WA, Drier Y, Liau BB, et al. Insulator dysfunction and oncogene activation in IDH mutant gliomas. Nature 2016; 529:110–114.
8. Guo Y, Xu Q, Canzio D, et al. CRISPR inversion of CTCF sites alters genome topology and enhancer/promoter function. Cell 2015; 162:900–910.
9. Heinz S, Romanoski CE, Benner C, et al. The selection and function of cell type-specific enhancers. Nat Rev Mol Cell Biol 2015; 16:144–154.
10. Pope SD, Medzhitov R. Emerging principles of gene expression programs and their regulation. Mol Cell 2018; 71:389–397.
11. Hnisz D, Schuijers J, Lin CY, et al. Convergence of developmental and oncogenic signaling pathways at transcriptional super-enhancers. Mol Cell 2015; 58:362–370.
12. Alder O, Cullum R, Lee S, et al. Hippo signaling influences HNF4A and FOXA2 enhancer switching during hepatocyte differentiation. Cell Rep 2014; 9:261–271.
13. Trompouki E, Bowman TV, Lawton LN, et al. Lineage regulators direct BMP and Wnt pathways to cell-specific programs during differentiation and regeneration. Cell 2011; 147:577–589.
14. Mullen AC, Orlando DA, Newman JJ, et al. Master transcription factors determine cell-type-specific responses to TGF-beta signaling. Cell 2011; 147:565–576.
15. Noble D. Conrad Waddington and the origin of epigenetics. J Exp Biol 2015; 218:816–818.
16. Vierbuchen T, Ling E, Cowley CJ, et al. AP-1 transcription factors and the BAF complex mediate signal-dependent enhancer selection. Mol Cell 2017; 68:1067–1082.e12.
17. Tessarz P, Kouzarides T. Histone core modifications regulating nucleosome structure and dynamics. Nat Rev Mol Cell Biol 2014; 15:703–708.
18. Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 2006; 126:663–676.
19. Giroux V, Rustgi AK. Metaplasia: tissue injury adaptation and a precursor to the dysplasia-cancer sequence. Nat Rev Cancer 2017; 17:594–604.
20. Kopp JL, Grompe M, Sander M. Stem cells versus plasticity in liver and pancreas regeneration. Nat Cell Biol 2016; 18:238–245.
21. Zhou Q, Melton DA. Pancreas regeneration. Nature 2018; 557:351–358.
22. Beumer J, Clevers H. Regulation and plasticity of intestinal stem cells during homeostasis and regeneration. Development 2016; 143:3639–3649.
23. Gonzales KAU, Fuchs E. Skin and its regenerative powers: an alliance between stem cells and their niche. Dev Cell 2017; 43:387–401.
24. Dor Y, Brown J, Martinez OI, et al. Adult pancreatic beta-cells are formed by self-duplication rather than stem-cell differentiation. Nature 2004; 429:41–46.
25. Blanpain C, Simons BD. Unravelling stem cell dynamics by lineage tracing. Nat Rev Mol Cell Biol 2013; 14:489–502.
26. Dhawan S, Georgia S, Tschen SI, et al. Pancreatic beta cell identity is maintained by DNA methylation-mediated repression of Arx. Dev Cell 2011; 20:419–429.
27. Papizan JB, Singer RA, Tschen SI, et al. Nkx2.2 repressor complex regulates islet beta-cell specification and prevents beta-to-alpha-cell reprogramming. Genes Dev 2011; 25:2291–2305.
28. Chakravarthy H, Gu X, Enge M, et al. Converting adult pancreatic islet alpha cells into beta cells by targeting both Dnmt1 and Arx. Cell Metab 2017; 25:622–634.
29. Li J, Casteels T, Frogne T, et al. Artemisinins target GABAA receptor signaling and impair alpha cell identity. Cell 2017; 168:86–100.e15.
30. Ben-Othman N, Vieira A, Courtney M, et al. Long-term GABA administration induces alpha cell-mediated beta-like cell neogenesis. Cell 2017; 168:73–85.e11.
31. Thorel F, Népote V, Avril I, et al. Conversion of adult pancreatic alpha-cells to beta-cells after extreme beta-cell loss. Nature 2010; 464:1149–1154.
32. Chera S, Baronnier D, Ghila L, et al. Diabetes recovery by age-dependent conversion of pancreatic delta-cells into insulin producers. Nature 2014; 514:503–507.
33. Collombat P, Xu X, Ravassard P, et al. The ectopic expression of Pax4 in the mouse pancreas converts progenitor cells into alpha and subsequently beta cells. Cell 2009; 138:449–462.
34. Yang YP, Thorel F, Boyer DF, et al. Context-specific alpha-to-beta-cell reprogramming by forced Pdx1 expression. Genes Dev 2011; 25:1680–1685.
35. Xiao X, Guo P, Shiota C, et al. Endogenous reprogramming of alpha cells into beta cells, induced by viral gene therapy, reverses autoimmune diabetes. Cell Stem Cell 2018; 22:78–90.e4.
36. Bernstein BE, Mikkelsen TS, Xie X, et al. A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell 2006; 125:315–326.
37. Bramswig NC, Everett LJ, Schug J, et al. Epigenomic plasticity enables human pancreatic alpha to beta cell reprogramming. J Clin Invest 2013; 123:1275–1284.
38. Wiles ET, Selker EU. H3K27 methylation: a promiscuous repressive chromatin mark. Curr Opin Genet Dev 2017; 43:31–37.
39. Neiman D, Moss J, Hecht M, et al. Islet cells share promoter hypomethylation independently of expression, but exhibit cell-type-specific methylation in enhancers. Proc Natl Acad Sci U S A 2017; 114:13525–13530.
40. Kloppel G. Neuroendocrine neoplasms: dichotomy, origin and classifications. Visc Med 2017; 33:324–330.
41. Hermann G, Konukiewitz B, Schmitt A, et al. Hormonally defined pancreatic and duodenal neuroendocrine tumors differ in their transcription factor signatures: expression of ISL1, PDX1, NGN3, and CDX2. Virchows Arch 2011; 459:147–154.
42. Jiao Y, Shi C, Edil BH, et al. DAXX/ATRX, MEN1, and mTOR pathway genes are frequently altered in pancreatic neuroendocrine tumors. Science 2011; 331:1199–1203.
43. Elsasser SJ, Allis CD, Lewis PW. Cancer. New epigenetic drivers of cancers. Science 2011; 331:1145–1146.
44. Larsson C, Skogseid B, Oberg K, et al. Multiple endocrine neoplasia type 1 gene maps to chromosome 11 and is lost in insulinoma. Nature 1988; 332:85–87.
45. Chandrasekharappa SC, Guru SC, Manickam P, et al. Positional cloning of the gene for multiple endocrine neoplasia-type 1. Science 1997; 276:404–407.
46. Lu J, Herrera PL, Carreira C, et al. Alpha cell-specific Men1 ablation triggers the transdifferentiation of glucagon-expressing cells and insulinoma development. Gastroenterology 2010; 138:1954–1965.
47. Li F, Su Y, Cheng Y, et al. Conditional deletion of Men1 in the pancreatic beta-cell leads to glucagon-expressing tumor development. Endocrinology 2015; 156:48–57.
48. Chan CS, Laddha SV, Lewis PW, et al. ATRX, DAXX or MEN1 mutant pancreatic neuroendocrine tumors are a distinct alpha-cell signature subgroup. Nat Commun 2018; 9:4158.
49. Balogh K, Racz K, Patocs A, et al. Menin and its interacting proteins: elucidation of menin function. Trends Endocrinol Metab 2006; 17:357–364.
50. Matkar S, Thiel A, Hua X. Menin: a scaffold protein that controls gene expression and cell signaling. Trends Biochem Sci 2013; 38:394–402.
51. Hughes CM, Rozenblatt-Rosen O, Milne TA, et al. Menin associates with a trithorax family histone methyltransferase complex and with the hoxc8 locus. Mol Cell 2004; 13:587–597.
52. Yokoyama A, Cleary ML. Menin critically links MLL proteins with LEDGF on cancer-associated target genes. Cancer Cell 2008; 14:36–46.
53. Malik R, Khan AP, Asangani IA, et al. Targeting the MLL complex in castration-resistant prostate cancer. Nat Med 2015; 21:344–352.
54. Karnik SK, Hughes CM, Gu X, et al. Menin regulates pancreatic islet growth by promoting histone methylation and expression of genes encoding p27Kip1 and p18INK4c. Proc Natl Acad Sci U S A 2005; 102:14659–14664.
55. Agarwal SK, Guru SC, Heppner C, et al. Menin interacts with the AP1 transcription factor JunD and represses JunD-activated transcription. Cell 1999; 96:143–152.
56. Huang J, Gurung B, Wan B, et al. The same pocket in menin binds both MLL and JUND but has opposite effects on transcription. Nature 2012; 482:542–546.
57. Kim H, Lee JE, Cho EJ, et al. Menin, a tumor suppressor, represses JunD-mediated transcriptional activity by association with an mSin3A-histone deacetylase complex. Cancer Res 2003; 63:6135–6139.
58. Feng Z, Ma J, Hua X. Epigenetic regulation by the menin pathway. Endocr Relat Cancer 2017; 24:T147–T159.
59. Cao Y, Liu R, Jiang X, et al. Nuclear-cytoplasmic shuttling of menin regulates nuclear translocation of {beta}-catenin. Mol Cell Biol 2009; 29:5477–5487.
60. Jiang X, Cao Y, Li F, et al. Targeting beta-catenin signaling for therapeutic intervention in MEN1-deficient pancreatic neuroendocrine tumours. Nat Commun 2014; 5:5809.
61. Gurung B, Feng Z, Iwamoto DV, et al. Menin epigenetically represses Hedgehog signaling in MEN1 tumor syndrome. Cancer Res 2013; 73:2650–2658.
62. Goldberg AD, Banaszynski LA, Noh KM, et al. Distinct factors control histone variant H3.3 localization at specific genomic regions. Cell 2010; 140:678–691.
63. Lewis PW, Elsaesser SJ, Noh KM, et al. Daxx is an H3.3-specific histone chaperone and cooperates with ATRX in replication-independent chromatin assembly at telomeres. Proc Natl Acad Sci U S A 2010; 107:14075–14080.
64. Dyer MA, Qadeer ZA, Valle-Garcia D, et al. ATRX and DAXX: mechanisms and mutations. Cold Spring Harb Perspect Med 2017; 7:1–16.
65. Heaphy CM, de Wilde RF, Jiao Y, et al. Altered telomeres in tumors with ATRX and DAXX mutations. Science 2011; 333:425.
66. Marinoni I, Kurrer AS, Vassella E, et al. Loss of DAXX and ATRX are associated with chromosome instability and reduced survival of patients with pancreatic neuroendocrine tumors. Gastroenterology 2014; 146:453–460.e5.
67. Danussi C, Bose P, Parthasarathy PT, et al. Atrx inactivation drives disease-defining phenotypes in glioma cells of origin through global epigenomic remodeling. Nat Commun 2018; 9:1057.
68. Aguayo-Mazzucato C, Bonner-Weir S. Pancreatic beta cell regeneration as a possible therapy for diabetes. Cell Metab 2018; 27:57–67.
69. Zare M, Rastegar S, Ebrahimi E, et al. Role of pancreatic duct cell in beta cell neogenesis: a mini review study. Diabetes Metab Syndr 2017; 11: (Suppl 1): S1–S4.
70. Inada A, Nienaber C, Katsuta H, et al. Carbonic anhydrase II-positive pancreatic cells are progenitors for both endocrine and exocrine pancreas after birth. Proc Natl Acad Sci U S A 2008; 105:19915–19919.
71. Lankisch PG, Apte M, Banks PA. Acute pancreatitis. Lancet 2015; 386:85–96.
72. Kleeff J, Whitcomb DC, Shimosegawa T, et al. Chronic pancreatitis. Nat Rev Dis Primers 2017; 3:17060.
73. De Groef S, Leuckx G, Van Gassen N, et al. Surgical injury to the mouse pancreas through ligation of the pancreatic duct as a model for endocrine and exocrine reprogramming and proliferation. J Vis Exp 2015; 1:e52765.
74. Yasuda H, Kataoka K, Ichimura H, et al. Cytokine expression and induction of acinar cell apoptosis after pancreatic duct ligation in mice. J Interferon Cytokine Res 1999; 19:637–644.
75. Van Gassen N, Van Overmeire E, Leuckx G, et al. Macrophage dynamics are regulated by local macrophage proliferation and monocyte recruitment in injured pancreas. Eur J Immunol 2015; 45:1482–1493.
76. Foster JR. A review of animal models of nonneoplastic pancreatic diseases. Toxicol Pathol 2014; 42:243–259.
77. Storz P. Acinar cell plasticity and development of pancreatic ductal adenocarcinoma. Nat Rev Gastroenterol Hepatol 2017; 14:296–304.
78. Ziv O, Glaser B, Dor Y. The plastic pancreas. Dev Cell 2013; 26:3–7.
79. Strobel O, Dor Y, Alsina J, et al. In vivo lineage tracing defines the role of acinar-to-ductal transdifferentiation in inflammatory ductal metaplasia. Gastroenterology 2007; 133:1999–2009.
80. Rooman I, Real FX. Pancreatic ductal adenocarcinoma and acinar cells: a matter of differentiation and development? Gut 2012; 61:449–458.
81. Pan FC, Bankaitis ED, Boyer D, et al. Spatiotemporal patterns of multipotentiality in Ptf1a-expressing cells during pancreas organogenesis and injury-induced facultative restoration. Development 2013; 140:751–764.
82. Kamisawa T, Wood LD, Itoi T, et al. Pancreatic cancer. Lancet 2016; 388:73–85.
83. Ryan DP, Hong TS, Bardeesy N. Pancreatic adenocarcinoma. N Engl J Med 2014; 371:1039–1049.
84. Makohon-Moore A, Iacobuzio-Donahue CA. Pancreatic cancer biology and genetics from an evolutionary perspective. Nat Rev Cancer 2016; 16:553–565.
85. Hruban RH, Adsay NV, Albores-Saavedra J, et al. Pancreatic intraepithelial neoplasia: a new nomenclature and classification system for pancreatic duct lesions. Am J Surg Pathol 2001; 25:579–586.
86. Pinho AV, Chantrill L, Rooman I. Chronic pancreatitis: a path to pancreatic cancer. Cancer Lett 2014; 345:203–209.
87. Guerra C, Schuhmacher AJ, Cañamero M, et al. Chronic pancreatitis is essential for induction of pancreatic ductal adenocarcinoma by K-Ras oncogenes in adult mice. Cancer Cell 2007; 11:291–302.
88. Morris JP 4th, Cano DA, Sekine S, et al. Beta-catenin blocks Kras-dependent reprogramming of acini into pancreatic cancer precursor lesions in mice. J Clin Invest 2010; 120:508–520.
89. Kopp JL, von Figura G, Mayes E, et al. Identification of Sox9-dependent acinar-to-ductal reprogramming as the principal mechanism for initiation of pancreatic ductal adenocarcinoma. Cancer Cell 2012; 22:737–750.
90. Habbe N, Shi G, Meguid RA, et al. Spontaneous induction of murine pancreatic intraepithelial neoplasia (mPanIN) by acinar cell targeting of oncogenic Kras in adult mice. Proc Natl Acad Sci U S A 2008; 105:18913–18918.
91. Ji B, Tsou L, Wang H, et al. Ras activity levels control the development of pancreatic diseases. Gastroenterology 2009; 137:1072–1082. 1082.e1–1082.e6.
92. Hingorani SR, Petricoin EF, Maitra A, et al. Preinvasive and invasive ductal pancreatic cancer and its early detection in the mouse. Cancer Cell 2003; 4:437–450.
93. Krapp A, Knöfler M, Frutiger S, et al. The p48 DNA-binding subunit of transcription factor PTF1 is a new exocrine pancreas-specific basic helix-loop-helix protein. EMBO J 1996; 15:4317–4329.
94. Krapp A, Knöfler M, Ledermann B, et al. The bHLH protein PTF1-p48 is essential for the formation of the exocrine and the correct spatial organization of the endocrine pancreas. Genes Dev 1998; 12:3752–3763.
95. Masui T, Swift GH, Deering T, et al. Replacement of Rbpj with Rbpjl in the PTF1 complex controls the final maturation of pancreatic acinar cells. Gastroenterology 2010; 139:270–280.
96. Pin CL, Rukstalis JM, Johnson C, et al. The bHLH transcription factor Mist1 is required to maintain exocrine pancreas cell organization and acinar cell identity. J Cell Biol 2001; 155:519–530.
97. Direnzo D, Hess DA, Damsz B, et al. Induced Mist1 expression promotes remodeling of mouse pancreatic acinar cells. Gastroenterology 2012; 143:469–480.
98. Lo HG, Jin RU, Sibbel G, et al. A single transcription factor is sufficient to induce and maintain secretory cell architecture. Genes Dev 2017; 31:154–171.
99. Holmstrom SR, Deering T, Swift GH, et al. LRH-1 and PTF1-L coregulate an exocrine pancreas-specific transcriptional network for digestive function. Genes Dev 2011; 25:1674–1679.
100. von Figura G, Morris JP 4th, Wright CV, et al. Nr5a2 maintains acinar cell differentiation and constrains oncogenic Kras-mediated pancreatic neoplastic initiation. Gut 2014; 63:656–664.
101. Cobo I, Martinelli P, Flández M, et al. Transcriptional regulation by NR5A2 links differentiation and inflammation in the pancreas. Nature 2018; 554:533–537.
102. Jiang M, Azevedo-Pouly AC, Deering TG, et al. MIST1 and PTF1 collaborate in feed-forward regulatory loops that maintain the pancreatic acinar phenotype in adult mice. Mol Cell Biol 2016; 36:2945–2955.
103. Hale MA, Swift GH, Hoang CQ, et al. The nuclear hormone receptor family member NR5A2 controls aspects of multipotent progenitor cell formation and acinar differentiation during pancreatic organogenesis. Development 2014; 141:3123–3133.
104. Zhu L, Tran T, Rukstalis JM, et al. Inhibition of Mist1 homodimer formation induces pancreatic acinar-to-ductal metaplasia. Mol Cell Biol 2004; 24:2673–2681.
105. Shi G, Zhu L, Sun Y, et al. Loss of the acinar-restricted transcription factor Mist1 accelerates Kras-induced pancreatic intraepithelial neoplasia. Gastroenterology 2009; 136:1368–1378.
106. Hoang CQ, Hale MA, Azevedo-Pouly AC, et al. Transcriptional maintenance of pancreatic acinar identity, differentiation, and homeostasis by PTF1A. Mol Cell Biol 2016; 36:3033–3047.
107. Flandez M, Cendrowski J, Cañamero M, et al. Nr5a2 heterozygosity sensitises to, and cooperates with, inflammation in KRas(G12V)-driven pancreatic tumourigenesis. Gut 2014; 63:647–655.
108. Seymour PA, Freude KK, Tran MN, et al. SOX9 is required for maintenance of the pancreatic progenitor cell pool. Proc Natl Acad Sci U S A 2007; 104:1865–1870.
109. Shih HP, Kopp JL, Sandhu M, et al. A Notch-dependent molecular circuitry initiates pancreatic endocrine and ductal cell differentiation. Development 2012; 139:2488–2499.
110. Wei D, Wang L, Yan Y, et al. KLF4 is essential for induction of cellular identity change and acinar-to-ductal reprogramming during early pancreatic carcinogenesis. Cancer Cell 2016; 29:324–338.
111. He P, Yang JW, Yang VW, et al. Kruppel-like factor 5, increased in pancreatic ductal adenocarcinoma, promotes proliferation, acinar-to-ductal metaplasia, pancreatic intraepithelial neoplasia, and tumor growth in mice. Gastroenterology 2018; 154:1494–1508.e13.
112. Das KK, Heeg S, Pitarresi JR, et al. ETV5 regulates ductal morphogenesis with Sox9 and is critical for regeneration from pancreatitis. Dev Dyn 2018; 247:854–866.
113. Miyatsuka T, Kaneto H, Shiraiwa T, et al. Persistent expression of PDX-1 in the pancreas causes acinar-to-ductal metaplasia through Stat3 activation. Genes Dev 2006; 20:1435–1440.
114. Roy N, Takeuchi KK, Ruggeri JM, et al. PDX1 dynamically regulates pancreatic ductal adenocarcinoma initiation and maintenance. Genes Dev 2016; 30:2669–2683.
115. Kubisch CH, Sans MD, Arumugam T, et al. Early activation of endoplasmic reticulum stress is associated with arginine-induced acute pancreatitis. Am J Physiol Gastrointest Liver Physiol 2006; 291:G238–G245.
116. Fazio EN, Young CC, Toma J, et al. Activating transcription factor 3 promotes loss of the acinar cell phenotype in response to cerulein-induced pancreatitis in mice. Mol Biol Cell 2017; 28:2347–2359.
117. Holmer R, Goumas FA, Waetzig GH, et al. Interleukin-6: a villain in the drama of pancreatic cancer development and progression. Hepatobiliary Pancreat Dis Int 2014; 13:371–380.
118. Corcoran RB, Contino G, Deshpande V, et al. STAT3 plays a critical role in KRAS-induced pancreatic tumorigenesis. Cancer Res 2011; 71:5020–5029.
119. Bailey JM, Hendley AM, Lafaro KJ, et al. p53 mutations cooperate with oncogenic Kras to promote adenocarcinoma from pancreatic ductal cells. Oncogene 2016; 35:4282–4288.
120. Lee AYL, Dubois CL, Sarai K, et al. Cell of origin affects tumour development and phenotype in pancreatic ductal adenocarcinoma. Gut 2018; 64:487–498.
121. Vakoc CR, Tuveson DA. Untangling the genetics from the epigenetics in pancreatic cancer metastasis. Nat Genet 2017; 49:323–324.
122. Boj SF, Hwang CI, Baker LA, et al. Organoid models of human and mouse ductal pancreatic cancer. Cell 2015; 160:324–338.
123. Roe JS, Hwang CI, Somerville TDD, et al. Enhancer reprogramming promotes pancreatic cancer metastasis. Cell 2017; 170:875–888.e20.
124. Mueller S, Engleitner T, Maresch R, et al. Evolutionary routes and KRAS dosage define pancreatic cancer phenotypes. Nature 2018; 554:62–68.
125. Diaferia GR, Balestrieri C, Prosperini E, et al. Dissection of transcriptional and cis-regulatory control of differentiation in human pancreatic cancer. EMBO J 2016; 35:595–617.
126. David CJ, Huang YH, Chen M, et al. TGF-beta tumor suppression through a lethal EMT. Cell 2016; 164:1015–1030.
127. Takaku M, Grimm SA, Wade PA. GATA3 in breast cancer: tumor suppressor or oncogene? Gene Expr 2015; 16:163–168.
128. Winslow MM, Dayton TL, Verhaak RG, et al. Suppression of lung adenocarcinoma progression by Nkx2-1. Nature 2011; 473:101–104.
129. Bailey P, Chang DK, Nones K, et al. Genomic analyses identify molecular subtypes of pancreatic cancer. Nature 2016; 531:47–52.
130. Andricovich J, Perkail S, Kai Y, et al. Loss of KDM6A activates super-enhancers to induce gender-specific squamous-like pancreatic cancer and confers sensitivity to BET inhibitors. Cancer Cell 2018; 33:512–526.e8.
131. Somerville TDD, Xu Y, Miyabayashi K, et al. TP63-mediated enhancer reprogramming drives the squamous subtype of pancreatic ductal adenocarcinoma. Cell Rep 2018; 25:1741–1755.e7.
132. St Pierre R, Kadoch C. Mammalian SWI/SNF complexes in cancer: emerging therapeutic opportunities. Curr Opin Genet Dev 2017; 42:56–67.
133. Kadoch C, Crabtree GR. Mammalian SWI/SNF chromatin remodeling complexes and cancer: mechanistic insights gained from human genomics. Sci Adv 2015; 1:e1500447.
134. Kadoch C, Hargreaves DC, Hodges C, et al. Proteomic and bioinformatic analysis of mammalian SWI/SNF complexes identifies extensive roles in human malignancy. Nat Genet 2013; 45:592–601.
135. Cancer Genome Atlas Research Network. Electronic address: [email protected]; Cancer Genome Atlas Research NetworkIntegrated genomic characterization of pancreatic ductal adenocarcinoma. Cancer Cell 2017; 32:185–203.e13.
136. Waddell N, Pajic M, Patch AM, et al. Whole genomes redefine the mutational landscape of pancreatic cancer. Nature 2015; 518:495–501.
137. von Figura G, Fukuda A, Roy N, et al. The chromatin regulator Brg1 suppresses formation of intraductal papillary mucinous neoplasm and pancreatic ductal adenocarcinoma. Nat Cell Biol 2014; 16:255–267.
138. Roy N, Malik S, Villanueva KE, et al. Brg1 promotes both tumor-suppressive and oncogenic activities at distinct stages of pancreatic cancer formation. Genes Dev 2015; 29:658–671.
139. Wang SC, Nassour I, Xiao S, et al. SWI/SNF component ARID1A restrains pancreatic neoplasia formation. Gut 2018; 68:1259–1270.
140. Kimura Y, Fukuda A, Ogawa S, et al. ARID1A maintains differentiation of pancreatic ductal cells and inhibits development of pancreatic ductal adenocarcinoma in mice. Gastroenterology 2018; 155:194–209.e2.
141. Wang W, Friedland SC, Guo B, et al. ARID1A, a SWI/SNF subunit, is critical to acinar cell homeostasis and regeneration and is a barrier to transformation and epithelial-mesenchymal transition in the pancreas. Gut 2018; 68:1245–1258.
142. Livshits G, Alonso-Curbelo D, Morris JP 4th, et al. Arid1a restrains Kras-dependent changes in acinar cell identity. Elife 2018; 7:1–28.
143. Ho L, Crabtree GR. Chromatin remodelling during development. Nature 2010; 463:474–484.
144. Dow LE, Lowe SW. Life in the fast lane: mammalian disease models in the genomics era. Cell 2012; 148:1099–1109.
145. Ventura A, Kirsch DG, McLaughlin ME, et al. Restoration of p53 function leads to tumour regression in vivo. Nature 2007; 445:661–665.
146. Dow LE, O’Rourke KP, Simon J, et al. Apc restoration promotes cellular differentiation and reestablishes crypt homeostasis in colorectal cancer. Cell 2015; 161:1539–1552.
Keywords:

Acinar-to-ductal metaplasia, Epigenetics, Pancreatic ductal adenocarcinoma, Pancreatic neuroendocrine tumor, Plasticitiy, Transcription

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