Novel Roles for the Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 in Development and Disease

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Citation Pearson, Daniel S. 2018. Novel Roles for the Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 in Development and Disease. Doctoral dissertation, Harvard University, Graduate School of Arts & Sciences.

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Novel Roles for the Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 in Development

and Disease

A dissertation presented

Daniel S. Pearson

The Division of Medical Sciences

in partial fulfillment of the requirements

for the degree of

Doctor of Philosophy

in the subject of

Biological and Biomedical Sciences

Harvard University

Cambridge, Massachusetts

March 2018

Dissertation Advisor: Professor George Q. Daley Daniel S. Pearson

Novel Roles for the Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 in Development

and Disease

Abstract

Terminal RNA uridylyltransferases (TUTases) are a class of non-canonical polyadenylate polymerases that are thought to function primarily by adding non-templated uridines to the 3’ ends of a wide range of RNA targets. Recently, two TUTases, ZCCHC6 and ZCCHC11, were shown to uridylate mRNAs, thereby marking them for degradation. While RNA uridylation is emerging as a potent mechanism of post-transcriptional gene regulation, uridylation-independent functions of

TUTases have also been reported. Despite these insights, the biological importance of

ZCCHC6/11 in physiologic and pathologic settings remains poorly understood. This dissertation explores the role of these TUTases in normal development and in cancer.

First, we examine Zcchc6/11 function in development and differentiation by profiling TUTase expression during mouse embryogenesis. In many tissues, we observe robust TUTase expression that decreases across developmental time, indicating a potential regulatory role. Using in vitro models of murine myogenesis, we find that Zcchc6/11 are regulators of muscle differentiation. Surprisingly, mutational analysis of Zcchc6 reveals that its catalytic activity is dispensable, while a portion of its N-terminal domain is required for this effect, indicating uridylation-independent mechanisms. Similarly, in mouse embryonic stem cells, TUTase downregulation contributes to the exit from naïve pluripotency, suggesting a broader regulatory

iii role in cellular differentiation. Collectively, these data indicate that TUTases regulate differentiation across diverse tissues and developmental stages.

Next, we investigate the impact of ZCCHC6/11 in cancer. Expression profiling reveals low levels of the TUTases in most normal adult tissues, yet robust expression across a variety of cancer cell lines and primary tumors. Consistent with an oncofetal pattern of expression, many adult tissues where TUTase expression is absent also exhibit reactivation to fetal levels following malignant transformation. Functionally, we demonstrate that ZCCHC6 supports the growth and tumorigenicity of a subset of cancer cell lines. Mechanistically, these effects are associated with altered mRNA turnover, including dysregulation of cell cycle factors and histone proteins. Finally, we find that CRISPR-Cas9 mediated knockout of ZCCHC6 sensitizes cancer cells to DNA damaging agents, potentially through the disruption of histone turnover. Overall, our results uncover novel functional roles for ZCCHC6/11, with therapeutic implications for both tissue regeneration and cancer.

TABLE OF CONTENTS

Acknowledgements ...... vii List of Figures ...... ix Chapter 1: Background ...... 1 1.1.Introduction ...... 2 Post-transcriptional gene regulation: a brief overview ...... 4 1.2. The Epitranscriptome ...... 6 mRNA polyadenylation ...... 6 1.3. TUTases: a class of non-canonical poly(A) polymerases ...... 8 Cid1: the prototypic TUTase ...... 10 1.4. Terminal RNA uridylation and gene regulation ...... 10 Histone mRNA uridylation ...... 10 Poly(A) mRNA uridylation ...... 13 Uridylation and miRNA biogenesis ...... 15 Mature miRNA uridylation...... 18 Uridylation as a quality control mechanism ...... 19 Uridylation-independent functions of the TUTases ...... 20 1.5. Known biologic functions of TUTases ...... 21 Chapter 2: The Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 Regulate Stem and Progenitor Cell Fate in a Catalytic-independent Manner ...... 23 2.1. Introduction ...... 24 2.2. Results ...... 26 TUTases are developmentally regulated genes ...... 26 TUTases contribute to primary myoblast differentiation………………………………………. 28 TUTase regulation of myogenesis is uridylation-independent ...... 31 TUTase overexpression blunts the induction of differentiation genes ...... 35 TUTase overexpression modestly impacts mRNA half-lives ...... 39 An N-terminal region of Zcchc6 is required for the regulation of myogenesis...... 40 TUTases regulate the exit from naïve pluripotency ...... 41 2.3. Discussion ...... 45 2.4. Methods ...... 48 Chapter 3: The Role of the Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 in Cancer ...... 58 3.1. Introduction ...... 59

3.2 Results ...... 60 TUTases exibit an oncofetal pattern of expression ...... 60 ZCCHC6/11 are dispensable for NIH3T3 transformation by KRASG12V ...... 63 ZCCHC6 supports the growth and tumorigenicity in a subset of cancer cell lines ...... 64 ZCCHC6 regulates mRNA uridylation and turnover in cancer cells ...... 67 ZCCHC6 impacts the cell cycle in cancer cells ...... 69 ZCCHC6 depletion sensitizes cancer cells to DNA damage ...... 71 ZCCHC6/11 are dispensable for tumorigenesis in the APCmin model of colon cancer ...... 73 3.3. Discussion ...... 75 3.4. Methods ...... 77 Chapter 4: Discussion and Perspective ...... 87 4.1. TUTases are regulators of cell state transitions...... 88 4.2. Uridylation-dependent vs. independent functions of the TUTases...... 90 4.3. Regulation of the cell cycle: a unifying theme? ...... 92 4.4. Therapeutic implications of TUTase modulation ...... 93 Appendix: Supplementary Material ...... 95 Bibliography ...... 107

ACKNOWLEDGEMENTS

I am extraordinarily grateful for the support and mentorship of Dr. George Q. Daley over the last several years. By providing critical feedback, enduring enthusiasm in the face of numerous failures, and the freedom to pursue scientific questions that I find interesting, he created a graduate experience that stimulated tremendous growth, both scientific and personal.

I would also like to thank my many previous mentors: Drs. Francis Collins, Michael Stitzel,

Praveen Sethupathy and Pramod Mistry for fostering my scientific curiosity and providing the opportunities that led me to pursue graduate training. Their continuing support and encouragement have been invaluable.

Scientific inquiry doesn’t happen in isolation and my experience in the Daley laboratory has been shaped by many wonderful colleagues. In particular, I would like to thank Dr. Kaloyan

Tsanov, a senior graduate student whom I worked very closely with over the major of my graduate experience. Through the highs and lows, Kal’s perpetual positive attitude and infectious passion for molecular biology provided incredible emotional and technical support over the years and resulted in what I anticipate to be an enduring friendship.

I would also like to thank my other close collaborators in the lab for their contributions to the projects described in this dissertation: Drs. Yu-Chung “Harry” Huang, Areum Haan, and Pavlos

Missios. I feel very fortunate having had the opportunity work with this talented group of scientists.

Their generosity came across countless times with their willingness to read and give critical feedback on data, paper drafts and committee meeting reports. Additionally, I am grateful for Dr.

John Powers’ creativity, optimism, and advice over the years.

I would also like to thank the technicians with whom I have worked with: Jess Barragan and

Grace LaPier. In addition, thank you the many present and past members of the Daley Lab including: Jihan Osborne, Alena Yermalovich, Deepak Jha, Ho-Chou Tu Sutheera “Mam”

Ratanasirintrawoot, Marcella Cesana, Sergei Doulatov, Linda Vo, Sam Morris, Patrick Cahan,

Aimee Dixon, Kathryn Entner, Judit Totth, Beth Kaleta, Patricia Sousa and Willy Lensch. In

vii particular, to our current lab manager, Michael Morse, for his many wild ideas and unwavering enthusiasm for science.

Thank you to my Dissertation Advisory Committee, Drs. John Rinn, Stephen Buratowski, and

Robert Horvitz for their feedback and scientific guidance over the years. For the contributions of collaborators from other labs including Drs. Oliver Hofmann, Lorena Pantano, Matthew Jones, and Dónal O’Carroll.

I would like to thank the BBS Program for their support both programmatic and personal over the years, Trista North for volunteering to read and provide critical feedback on this dissertation, and to my friends outside of the lab especially to Vipul Kumar for his wisdom and encouragement.

Finally, I would like to thank my family. My parents, Matthew, Holly and my wife Laurie for their love and unwavering support. I am here today because of you.

viii

LIST OF FIGURES

Figure 1.1. The “Central Dogma” of molecular biology and the epitranscriptome……………….3 Figure 1.2. Protein domain organization of human TUTases………………………………………9 Figure 1.3. Uridylation regulates histone and poly(A) mRNA fate ...... 11 Figure 1.4. Uridylation regulates microRNA biogenesis and function ...... 16 Figure 2.1. TUTases are developmentally regulated genes ...... 27 Figure 2.2. TUTases contribute to primary myoblast differentiation ...... 29 Figure 2.3. TUTase regulation of myogenesis is uridylation-independent…….. . ………………33 Figure 2.4. TUTase overexpression blunts the induction of differentiation genes ...... 36 Figure 2.5. The N-terminus of Zcchc6 is required for the regulation of myogenesis ...... 40 Figure 2.6. TUTases contribute to the exit from naïve pluripotency ...... 42 Figure 3.1. TUTases exhibit an oncofetal pattern of expression…………………………...... 61 Figure 3.2. ZCCHC6 supports the growth and tumorigenicity in a subset of cancer cell lines . 66 Figure 3.3. ZCCHC6 regulates mRNA uridylation and turnover in cancer cell………………….68 Figure 3.4. ZCCHC6 depletion disrupts the cell cycle……………………………………………. 70 Figure 3.5. ZCCHC6 depletion sensitizes cancer cells to DNA damage…………………….. ... 72

CHAPTER 1

Background

1.1. Introduction.

Humans are multicellular organisms composed of roughly 37 trillion cells comprised of at least

200 morphologically distinct cell types which differ tremendously in terms of size, form and function1,2. With advances in single-cell transcriptomic analysis, the number of distinct cell types as defined by gene expression will certainly rise by orders of magnitude3. Though the characteristics that define a particular cell type is a matter of ongoing debate, it is well accepted that variation in gene expression, rather than differences in DNA content, gives rise to a particular cell type4. Many important processes in development and disease are driven by dramatic changes in gene expression. Therefore, understanding the gene regulatory programs orchestrating organismal developmental, as well as the contribution of dysregulated gene expression to disease etiology, have been areas of intense investigation in recent decades.

The control of gene expression, at first glance, is hugely complex as there are numerous layers of regulation. However, an organizing principle termed “The Central Dogma” of molecular biology, described by Francis Crick nearly 60 years ago5, is still used as a framework for describing much of gene regulation as it is understood today. This principle posits that genetic information, encoded within genomic DNA, is carried by messenger molecules which then dictate the expression of specific proteins (Figure 1.1A). Those carrier molecules were later identified as messenger RNAs (mRNAs)6. While mRNAs, and other classes of non-coding RNAs (ncRNAs), have critical functions beyond protein coding7,8, mRNAs and their protein coding function have remained a central focus of gene regulation.

mRNA expression is dynamically controlled at many stages throughout its lifespan, from their origins during transcription to the regulation of their decay. The nascent pre-mRNA transcribed by RNA Polymerase II is extensively modified both co-transcriptionally and after transcriptional termination giving rise to a mature mRNA molecule9. Mature mRNAs are exported to the cytoplasm where they direct the translation of their encoded protein.

Figure 1.1. The “Central Dogma” of molecular biology and the epitranscriptome. (A) Schematic of the “Central Dogma” of molecular biology. (B) Overview of frequently occurring epitranscriptomic modifications of mRNAs. Chemical structures of modified ribonucleosides are shown with altered residues highlighted in red. For 3’ end modifications text size represents relative frequency of each tail length. Frequencies of 3’ end modifications were obtained from Chang et al. 2014, Molecular Cell. Adapted from Xiaoyu et al. 2016, Nature Methods. Note, post-transcriptional gene regulation (PGTR) and RNA binding proteins (RBPs).

Historically, transcriptional regulation of mRNAs has been one of the most heavily studied aspects of eukaryotic gene regulation10. Transcription factors (TFs) are a major class of regulators that play important roles in controlling both the initiation and overall level of transcription11. TF expression is often restricted to certain tissues. Thus, TFs can play critical roles as master regulators of cell identity by driving tissue specific gene expression. For example,

3 ectopic overexpression of a single muscle-specific transcription factor, Myogenic Differentiation 1

(MyoD), is sufficient to convert fibroblasts into myoblasts12,13. Similarly, in cancer, disruption of transcriptional regulation is a core feature of most malignancies14 and aberrant overexpression of certain transcription factors, such as c-MYC, play critical roles in oncogenesis15. While certainly important, most transcription factors are not “master regulators” and their transcriptional regulation must work in concert with other aspects of gene regulatory control to finely tune both mRNA and protein expression specific to a select cell type or function16.

Post-transcriptional gene regulation: a brief overview

Post-transcriptional gene regulation (PTGR) has more recently emerged as a potent layer of regulatory control and, in certain cellular contexts such as the very early stages of embryonic development, it is the dominate form of gene regulation17,18. Furthermore, global transcriptional and proteomic analyses across diverse cell types, from embryonic stem cells to cancer cell lines, have revealed relatively poor correlations between mRNA and protein abundance, indicating that there must be prominent PTGR mechanisms at play19–22. Collectively, PTGR mechanisms cooperate with transcriptional control to shape the transcriptome, the magnitude of protein expression, the spatial and temporal aspects of translational initiation, and localization of mRNAs.

These actions are mainly carried out by two large classes of regulators, ncRNAs and RNA binding proteins (RBPs).

MicroRNAs (miRNAs) are small ncRNAs that are one of the most extensively characterized effectors of PTGR. miRNAs negatively regulate specific mRNA targets by impairing translation or stimulating mRNA decay23. Similar to TFs, miRNA expression is often tissue specific24 and miRNAs are known to be important regulators in both development and disease25. Further highlighting their importance, classification by miRNA expression profiling outperformed mRNA profiling at identifying poorly differentiated tumors across a wide range of human cancers26.

In humans, there are estimated to be well over 1000 RNA-binding proteins (RBPs)27. These core effectors of PTGR influence RNA localization, stability and mRNA translation through direct interactions with RNA resulting in the formation of ribonucleoprotein complexes (RNPs). For example, miRNAs exert their function through the RNP RNA-induced silencer complex (RISC)28.

In addition, RBPs can directly bind mRNAs and modulate the accessibility of miRNA target sites on mRNAs, thus sparing or sensitizing specific mRNAs to miRNA-mediated repression29.

A major function of RBPs is the direct regulation of mRNA stability, which in turn can control mRNA abundance as well as the association with translational machinery30. Additionally, RBPs are essential machinery in mRNA decay pathways and can contribute to quality control mechanisms that identify and clear aberrantly synthesized or damaged RNAs31. The regulation of mRNA stability often involves a combination of sequence specific elements intrinsic to certain mRNAs, such as AU-rich elements (AREs)32, as well as additional features that are discussed later including mRNA polyadenylate (poly(A)) tail length. AREs are one of the best characterized sequence elements. They are recognized by a range of RBPs33 and are generally found on mRNAs whose stability is quite labile, including many cell cycle regulators, cytokines and transcription factors34. Depending on the identity of the RBP, the target mRNA will either be stabilized or targeted for degradation, which can influence cell fate decisions. For example, human antigen R (HuR) promotes muscle differentiation through preferentially stabilizing mRNAs of critical transcriptional regulators of myogenesis including MyoD, Myogenin and p2135. In contrast, AU-binding factor 1 (AUF1)-mediated destabilization of mRNA targets is critical for muscle stem cell function during regeneration in response to injury36.

Lastly, a group of RBPs can chemically modify a wide range of RNA targets including mRNAs.

The totality of RNAs present in a cell or a cell population is called the transcriptome. Therefore, the study of these dynamic RNA modifications is referred to as “RNA epigenetics37” or

“epitranscriptomics38” meaning “above the transcriptome.” Much less is known about epitranscriptional modifications relative to the other mechanisms of PTGR discussed above.

However, emerging evidence indicates these modifications have the potential to influence gene expression, thus implicating their likely importance to physiologic and pathologic processes.

1.2. The Epitranscriptome.

The epitranscriptome consists of two major classes of epitranscriptomic marks, chemical modifications of RNA nucleosides and non-templated terminal 3’ base additions39,40. There are more than 100 known chemical modifications of RNA nucleosides that have been identified to- date including: N6-methyladenosine (m6A), N1-methyladenosine (m1A), 5-methylcytidine (m5C), inosine, and pseudouridine. Some modifications, such as pseudouridine, were first identified decades ago and were thought to occur primarily in transfer RNAs (tRNAs) and other non-coding

RNAs (ncRNAs)41. Due to advances in unbiased transcriptome-wide profiling techniques, it is now known that many epitranscriptomic modifications occur frequently across virtually every class of

RNA including mRNAs39 (Figure 1.1B). Though the function of most epitranscriptomic modifications remains enigmatic, compelling case-studies indicate their importance to the regulation of mRNA stability42–44, localization45, and translation46,47. The biologic importance of epitranscriptomic base modifications and their emerging contributions to gene regulation has been reviewed in detail elsewhere48–50. I will instead focus on the second class of epitranscriptomic modifications: 3’ end modifications and review their enzymatic “writers”, known roles in gene regulation, and biologic functions.

mRNA Polyadenylation

In the early 1970s, it was discovered that mRNAs are polyadenylated post- transcriptionally51,52. Since then, there has been an accumulating wealth of data indicating that the poly(A) length influences numerous aspects of mRNA metabolism and function53. The 3’ poly(A) tail is a nearly universal feature of mRNAs and is thus, by far, the most common 3’ end modification of mRNAs.

mRNAs are polyadenylated immediately after transcriptional termination with initial poly(A) tail lengths being upwards of 200 nucleotides (nt). Tail length is determined by both DNA sequence motifs and trans-acting proteins. The vast majority of mRNAs have a poly(A) signal (PAS), which contains an (AAUAAA) motif that is located ~10-30nt upstream of the cleavage/polyadenylation site, as well as a second GU-rich motif located ~20-40nt downstream of the cleavage/polyadenylation site54. The sequence composition of this second site, as well as the distance from the PAS, determines the location of nascent strand cleavage and initial poly(A) tail length55,56. A large multi-protein complex containing multiple cleavage factors and the canonical

RNA polyadenylate polymerase (PAP) recognizes these two motifs and catalyzes cleavage and subsequent polyadenylation57. Poly(A)-binding protein II (PAB II) also influences initial poly(A) tail length by enhancing PAP polyadenylation through stabilization of the PAP elongation complex via protein-protein interactions with PAP58. Finally, polyadenylation is terminated once the elongation complex is destabilized59.

Changes in polyA tail length have been suggested to regulate both translation and RNA stability60,61. While there is some controversy surrounding the link between translational rates and poly(A) tail length62, there is abundant evidence linking mRNA stability to poly(A) tail length.

Deadenylation is a highly regulated process. Once in the cytoplasm, mRNA poly(A) tails are shortened by a number of deadenylases complexes. For many eukaryotic mRNAs, deadenylation is both the initiating and rate-limiting step of mRNA-degradation63. Most of what is known about mRNA deadenylation was first uncovered in yeast. The regulation of deadenylation is less well understood in humans, but given the high degree of conservation of deadenylases and exonucleases across species, similar mechanisms are likely at play64.

Deadenylation is usually a two-step process in which the PolyA Nucleases 2 and 3 (PAN2-

PAN3) first shortens poly(A) tails to around 110nts followed by additional shortening by the CCR4-

CAF1 poly(A) nuclease complex65. Once the poly(A) tail reaches a critical length, the decapping proteins 1 and 2 (DCP1/2) are recruited to remove the 5’ 7-methylguanosine cap resulting in the

7 initiation of 5’ to 3’ decay carried out by Exonuclease 1 (XRN1). Alternatively, degradation can continue in the 3’ to 5’ direction driven by a large multi-protein complex called the exosome66.

Additional deadenylases, such as Poly(A)-Specific Ribonuclease (PARN), can contribute to general RNA decay, but may have more specialized functions that are required in the context of

ARE-stimulated degradation67. Deadenylation is stimulated or perturbed by many pathways, including miRNA-mediated mRNA repression, as a means of controlling mRNA stability68.

Until the late 1980s, it was thought that the poly(A) tails of mRNAs were progressively shortened until RNA-decay was inevitably initiated. However, with the discovery of cytoplasmic adenylation, it became clear that mRNA poly(A) tail length is the result of dynamic balance between adenylation and deadenylation69. In addition to the elongation of poly(A) tails, candidate- based and transcriptome-wide studies revealed that non-templated 3’ adenines are a prominent feature of many classes of RNAs70–75. Surprisingly, these same studies also identified widespread non-templated 3’ uridines highlighting the importance of an enigmatic class of non-canonical

PAPs (ncPAPs) capable of adenylating or uridylating a wide range of RNAs.

1.3. TUTases: a class of non-canonical poly(A) polymerases.

Non-canonical PAPs are RBPs that are part of the larger DNA pol β superfamily76 and are conserved from yeast to humans70. In contrast to the canonical PAPs discussed above, ncPAPs function by adding non-templated nucleotides to the 3’ ends of a diverse array of RNA molecules, including polyadenylated mRNAs. These enzymes have the capacity to add non-templated adenines, uridines, and at lower frequencies, guanines and cytosines. A subset of ncPAPs, which favor uridines, are often referred to as terminal uridylyltransferases (TUTases)77. In humans, there are seven known TUTases (Figure 1.2). Emerging evidence suggests that these enzymes regulate gene expression through their nucleotide transferase activity via a variety of mechanisms, from modulation of mRNA stability to the regulation of miRNA biogenesis and function. All TUTases share a conserved nucleotidyl transferase domain, which contains the core

Figure 1.2. Protein domain organization of human TUTases. Domain structures of seven human TUTases and the S. pombe TUTase ortholog Cid1 aligned by their active catalytic domain. Domain identity is indicated by color according to the legend in the upper left corner. Protein sizes are indicated on the right. Note, AA = amino acid and PAP = poly(A) polymerase. Adapted from Norbury, C. J. 2013. Nat. Rev. Mol. Cell Biol. catalytic triad, and a poly(A) polymerase (PAP) associated domain which confers selectivity towards uridine nucleotides78. Structurally, among the TUTases, zinc finger CCHC domain- containing protein 6 (ZCCHC6) and zinc finger CCHC domain-containing protein 11 (ZCCHC11) stand out due to their large size and the presence of many additional domains including: multiple zinc fingers, one additional N-terminal nucleotidyl transferase, and PAP-associated domains.

Both of these N-terminal domains cannot contribute to catalysis because they harbor mutations that disrupt the catalytic triad and other critical residues responsible for uridine selectivity79.

Despite the minimal homology of ZCCHC6 and ZCCHC11 with the S. pombe ortholog Cid1, work over the last two decades have shown that the functions of these three genes are surprisingly similar.

Cid1: the prototypic TUTase

Cid1 (Caffeine induced death suppressor), one of the first TUTases to be discovered, was identified in S. pombe in a forward genetic screen for regulators of the DNA synthesis phase to mitosis (S-M) checkpoint80. Due to the structural similarity to known PAPs, Cid1 was initially thought to act as a cytoplasmic PAP which presumably polyadenylated members of the S-M checkpoint81. However, more refined biochemical characterization revealed that Cid1 was a

TUTase, as demonstrated by their remarkable preference for uridine relative to other nucleotides in reconstituted polymerization experiments70. Furthermore, upon S-phase cell cycle arrest, Cid1 uridylates a substantial fraction of actin mRNAs, which led to the presumption that this uridylation likely influences mRNA stability. It was later shown that Cid1 uridylates a large number of poly(A) mRNAs, and that terminal uridylation facilitates the binding of the Sm-like complex (LSM1-7) which recruits DCP1/2, thus initiating 5’ to 3’ RNA decay82. This suggested, for the first time, that mRNA uridylation might be a widespread and conserved mechanism to control poly(A) mRNA stability. From this point on, I will review the contributions of terminal uridylation to gene regulation, with a particular focus on uridylation directed by the TUTases ZCCHC6 and ZCCHC11.

1.4. Terminal RNA uridylation and gene regulation.

Histone mRNA uridylation

While accumulating evidence in Arabidopsis, mouse, and human systems indicated that uridylation might play a role in the decay of miRNA-mediated mRNA cleavage products83, the first compelling evidence that terminal uridylation contributed to the initiation of mRNA turnover came from the study of replication-dependent histone mRNA decay84. Eukaryotic genomic DNA is tightly packed in the nucleus and exists as a highly organized, yet dynamic, structure that resembles a fractal globule85. Proper regulation of DNA packaging is achieved in part due to four core histone proteins (H2A, H2B, H3 and H4), which form tetramers that dimerize to create an octamer in which

Figure 1.3. Uridylation regulates histone and poly(A) mRNA fate. (A) At the end of S-phase of the cell cycle, the multi-step process of histone mRNAs decay is promoted by terminal uridylation. Decay primarily continues in the 3’ to 5’ direction. In a minor parallel pathway, 5’ to 3’ decay can also contribute. (B) During mid-S-phase, 3’ ends of histone mRNAs are maintained by mono- and di-uridylation. (C) Terminal mRNA uridylation can promote decay and (D) inhibit protein translation. See text for a detailed explanation of each process. Adapted from Lee et al. 2014. Cell.

DNA is wound tightly around. Histone protein expression is highly regulated by multiple mechanisms, as imbalances between the abundance of histones and DNA lead to cytotoxicity86

11 and increased susceptibility to DNA damage87,88. Therefore, in the S-phase of the cell cycle, to account for the doubling of genomic DNA, histone protein levels must also be proportionally increased. To achieve this, histone mRNAs are only transcribed during DNA synthesis89.

Subsequently, these mRNAs are rapidly degraded at the end of S-phase90,91. Multiple mechanisms have been reported to regulate histone mRNA clearance, with terminal uridylation playing a prominent role in promoting decay.

Histone mRNAs differ from most other protein-coding mRNAs in that they lack a 3’ poly(A) tail. Instead, histone mRNAs have a highly conserved 25-26nt stem-loop structure near the 3’ terminus. This stem-loop is a major regulator that contributes to both the rapid transcription at the beginning of S-phase as well as the coordinated clearance of histone mRNAs at the end of S- phase90. To initiate decay, stem-loop binding protein (SLBP) binds the terminal stem-loop and, in concert with Up-Frameshift Suppressor 1 (UPF1), recruits a TUTase which uridylates the 3’ end of the histone mRNAs resulting in the initiation 3’ to 5’ degradation92. With the advent of high- coverage targeted RNA-sequencing, it became clear that uridylation is also a contributing factor to the progression of decay. It is now known that the stem loop is progressively degraded by 3’-

5’ Exoribonuclease 1 (ERI1) a few nucleotides at a time, followed by uridylation of the decay intermediate93,94. This back and forth between ERI1 and uridylation repeats until the whole stem loop is lost. Once the stem loop is degraded, the LSM1-7 complex is recruited and promotes 3’-

5’ decay in an exosome-dependent manner. Additionally, for a minority of decay products, terminal uridines can promote LSM1-7 binding, which stimulates decapping machinery to initiate

5’ to 3’ decay by XRN193 (Figure 1.3A).

The TUTase(s) responsible for oligouridylation across these stages of histone mRNA decay is a matter of some controversy. While there have been conflicting reports implicating multiple

TUTases92,95,96, a recent study provided compelling evidence that ZCCHC6 is the predominant

TUTase in uridylation-dependent histone mRNA decay97. Adding an additional layer of complexity to the impact of uridylation on histone mRNA stability, prior to the initiation of decay, mono and

12 di-uridylation may also be critical for maintaining the integrity of 3’ end, thus preventing decay prior to the end of S-phase98 (Figure 1.3B). Collectively, these studies confirmed that uridylation of mRNAs was not unique to yeast and indicated that mRNA uridylation might be a widely used and evolutionary conserved mechanism of PTGR.

Poly(A) mRNA uridylation Until recently, it was unclear if 3’ uridylation of poly(A) mRNAs occurs in mammals. This was primarily due to technical limitations of current high-throughput sequencing approaches. Standard

RNA-sequencing methods yield very poor coverage of 3’ ends due to biases introduced both during cDNA synthesis and template fragmentation prior to 3’ adaptor ligation99. Additionally, on commonly used sequencing platforms, accurate base calling near homopolymeric sequences, such as the poly(A) tail, is extremely challenging. Given these limitations, there is no evidence of terminal uridylation in standard RNA-seq data-sets. However, a transcriptome-wide approach called TAIL-seq recently overcame these technical hurdles by altering library construction to enrich for 3’ ends and creating a novel base calling algorithm well suited for homopolymeric stretches100.

TAIL-seq was originally developed as a transcriptome-wide method to accurately quantify poly(A) tail lengths. However, the authors noted that a substantial fraction of human and mouse mRNAs contain non-templated bases beyond their poly(A) tails with mono and di-uridylation being some of the most common additions. Additionally, they found 3’ uridylation was present most frequently on mRNAs with short poly(A) tails (<25 nt). There was also a correlation between frequently uridylated mRNAs and decreased mRNA half-life. Conversely, terminal guanines where found to be enriched on longer poly(A) tails, and guanylation of mRNAs correlated with longer half-lives suggesting an antagonistic role between terminal uridylation and guanylation.

Furthermore, upon transfection of mir-1 in HeLa cells, they observed an increase in the frequency

13 of mRNA uridylation in mir-1 targets, indicating that terminal mRNA uridylation may also play a role in miRNA-mediated silencing100.

Shortly after, ZCCHC6 and ZCCHC11 were shown to be the primary TUTases responsible for terminal mRNA uridylation in humans101. In addition, mRNA uridylation appears to be a global mechanism of influencing mRNA stability as siRNA depletion or CRISPR-Cas9-mediated knockout of either TUTase prolonged the half-lives of over 1000 mRNAs101. Combinatorial loss- of-function experiments further demonstrated that ZCCHC6 and ZCCHC11 act redundantly to regulate the stability of mRNAs through terminal uridylation. Furthermore, using a reconstituted mRNA uridylation assay, the addition of recombinant poly(A) binding protein (PABP) was shown to impair TUTase-directed terminal uridylation. This was likely mediated through direct competition between ZCCHC6/11 and PABP for binding to the poly(A) tail suggesting occupancy of RBPs on the poly(A) tail may shield certain mRNAs from uridylation-mediated decay. Finally, depletion of exonucleases involved in both 3’ to 5’ and 5’ to 3’ decay led to the accumulation of uridylated mRNA species, demonstrating that multiple pathways of mRNA clearance are regulated by terminal uridylation. While it remains unclear which pathways of RNA decay are most important in humans, the emerging model is quite similar to what has been shown for Cid1- directed mRNA decay in yeast (Figure 1.3C). Together, these data demonstrate that ZCCHC6/11 globally regulate mRNA stability through terminal uridylation in humans.

In addition to regulating the stability of mRNAs, the molecular consequences of uridylation may also depend on the cellular context and potentially varies between organisms. For example, in Xenopus oocytes, the ortholog of ZCCHC6, Xenopus Terminal Uridylyl Transferase 7 (XTUT7), was shown to direct mRNA oligouridylation of reporter gene mRNA transcripts, resulting in repression at the translational level without changes in mRNA abundance102. This is likely achieved by Watson-Crick base pairing between 3’ uridines added by TUTases and existing adenines of the poly(A) tail, resulting in a terminal hairpin. This structure may interfere with

14 translation by sterically hindering PABP binding to the poly(A) tail thereby disrupting the recruitment of translational initiation factors (Figure 1.3D). However, because mRNA decay pathways are known to be relatively quiescent in the oocyte103, it is unclear how generalizable this phenomenon might be. Taken together, these studies indicate that mRNA uridylation is a prevalent and evolutionarily conserved mechanism of gene regulation that has the potential to impact both mRNA stability and translation. However, ZCCHC6/11 directed terminal uridylation is not restricted to mRNAs.

Uridylation and miRNA biogenesis

MicroRNAs (miRNAs) are a highly conserved class of small (~22nt) single-stranded ncRNAs that have been implicated as critical negative regulators of mRNAs across numerous disease and developmental processes104. Global small RNA-sequencing data sets have revealed that the 3’ ends of both precursor and mature miRNAs are frequently modified with the vast majority of additions being short stretches (1-2nt) of uridines or adenines24,71–73,75. Those, and more recent studies, have demonstrated that miRNA-uridylation influences the biogenesis as well as the function of mature miRNAs through a range of mechanisms (Figure 1.4).

The vast majority of miRNAs are derived from RNA polymerase II-transcribed primary transcripts (pri-miRNA)105. These are recognized and processed by the DiGeorge Syndrome

Critical Region Gene 8 (DGCR8) and Double-Stranded RNA-Specific Endoribonuclease

(DROSHA) complex in the nucleus106. The resulting ~65nt hairpin precursor miRNA (pre-miRNA) is then transported to the cytoplasm by exportin-5106,107 and processed by DICER1 into a ~22nt double-stranded RNA duplex108–110. One of the RNA strands (guide-strand) is incorporated into the RNA-induced silencing complex (RISC)28. The guide-strand then guides RISC to specific mRNAs through Watson-Crick base-pairing resulting in translational inhibition and/or mRNA decay23.

Figure 1.4. Uridylation regulates microRNA biogenesis and function. Overview of terminal uridylation in microRNA biogenesis and function. 3’ non-templated uridylation of pre- and mature-miRNAs are shown in purple, while 3’ adenylation is shown in red. ZCCHC6 and ZCCHC11 are responsible for most of the pre- and mature-miRNA uridylation. See text for a detailed description of each function. Adapted from Thornton and Gregory, 2012. Trends in Cell Biology.

Cytoplasmic uridylation plays a major role in the regulation of pre-miRNAs processing by

DICER1, primarily by influencing the efficiency of this cleavage reaction. The majority of pre-

16 miRNAs (group I) are not uridylated because they have an optimal 2nt overhang on their 3’ ends making them a preferred substrate for DICER1 cleavage111. However, a second class of miRNAs (group II), which includes members of the let-7 family and mir-105-b, only have a 1nt 3’ overhang. The processing of group II miRNAs is impacted by ZCCHC6/11-dependent mono- uridylation that extends this overhang to 2nt resulting in enhanced DICER1 processing yielding increased mature miRNA expression77. A similar process serves as a quality-control surveillance system utilizing ZCCHC6/11 to identify pre-miRNAs that deviate more than 1nt from the optimal 3’ ends resulting in either end polishing to restore a 2nt overhang or, in the case of a severely resected 3’ end, oligouridylation followed by exosome-dependent 3’ to 5’ decay112,113.

In addition to globally regulating miRNA biogenesis, the TUTases are also involved in the targeted regulation of the let-7 family of miRNAs through a LIN28-dependent mechanism. LIN28A, and its paralog LIN28B, are highly conserved RNA binding proteins that regulate many processes including pluripotency, embryonic development, and cellular differentiation114. LIN28A/B exert their function through both direct mRNA binding115–117 and through the repression of the biogenesis of the let-7 family of miRNAs118–120. Mechanistically, LIN28A binds to pre-let-7 and recruits ZCCHC6/11, which leads to oligouridylation of pre-let7’s120–124. This polyuridine tail acts as a decay signal that is recognized by the uridylation-selective exonuclease DIS3 Like 3'-5'

Exoribonuclease 2 (DIS3L2), which degrades pre-let-7 in a 3’ to 5’ manner125–127. In cellular contexts where LIN28A or LIN28B are expressed, mature let-7 expression is usually quite low.

ZCCHC6/11 are thought to function redundantly in the LIN28/let-7 pathway and appear to be biochemically identical in vitro128. However, in vivo there are reports that ZCCHC11 is the dominant enzyme in determining the expression level of mature let-7 miRNAs125,128,129.

Mature miRNA uridylation

In addition to regulating miRNA biogenesis, mature miRNA function can also be altered by terminal uridylation. The binding of miRNA’s to their specific mRNA targets is primarily determined by the “seed” region of the miRNA (nucleotides 2-8 from the 5’)130. However, complementarity on the 3’ end of the miRNA can also contribute to mRNA target selection131. Thus, terminal uridylation of mature miRNAs could theoretically alter mRNA targeting through either disrupting or enhancing 3’ complementarity. While there are no documented cases of miRNA uridylation enhancing miRNA targeting, there are a number of instances where terminal uridylation has been shown to impair miRNA-directed mRNA repression. This phenomenon was first described in human and mouse cell lines where depletion of Zcchc11 impairs the production of a subset of cytokines in response to inflammatory stimuli72. For at least one of these cytokines, Interleukin 6

(IL-6), this effect was mechanistically linked to terminal uridylation of the mir-26 family. Normally,

IL-6 mRNA is modestly repressed by the mir-26 family. However, upon knockdown of Zcchc11,

IL-6 expression is strongly repressed. Strikingly, the vast majority of mir-26 reads contained at least one non-templated uridine. In contrast, mir-26 reads from Zcchc11 knockdown cells were virtually devoid of terminal uridylation. Reporter gene assays confirmed that terminal uridines can impair mir-26b mediated repression of IL-6. This is likely a widespread phenomenon, as it has also been reported for a number of miRNAs targeting Insulin Like Growth Factor 1 (IGF-1)132.

Certain miRNAs, such as the mir-26 family discussed above, are modified at much higher frequencies than others133. This raises the possibility that specificity factors, such as RBPs or motifs within mature miRNAs themselves, might preferentially recruit TUTases. Consistent with this hypothesis, a bipartite sequence motif present in some let-7 family members is both necessary and sufficient for Zcchc6/11 directed uridylation134. However, beyond this example, the mechanisms governing target specificity remain unclear.

Similar to the uridylation of poly(A) mRNAs, uridylation of mature miRNAs has also been reported to stimulate miRNA decay in the context of T-cell activation135. However, this effect on miRNA stability might be cell-type specific as most studies have not uncovered a causal relationship between miRNA abundance and terminal miRNA uridylation72,132,134,136. Interestingly, there have also been reports of 3’ adenylation of mature miRNAs that results in miRNA stabilization137–139. Thus, at least in some contexts, there are antagonistic effects of terminal uridylation and adenylation on both miRNAs and poly(A) mRNAs. In line with this observation, multiple studies have found that knockdown of the TUTases results in a decrease in the frequency of terminal miRNA uridylation, as well as a concomitant increase in the prevalence of terminal adenylation72,134.

The balance between mature miRNA uridylation and adenylation can also regulate miRNA localization. Exosomes are small membranous vesicles (30-100nm) that are derived from endosomic compartments and contain diverse molecular cargo, including proteins, mRNAs, and miRNAs140,141. These vesicles are secreted by most cell types and there is accumulating evidence in vivo that exosomes can travel to distant locations where their contents can contribute to gene regulatory role in their new location142. Small RNA-sequencing revealed that miRNAs in exosomes are uridylated at a much higher frequency than cytoplasmic miRNAs. While the

TUTases responsible are not yet known, these data indicate that miRNA incorporation into exosomes is potentially regulated by terminal uridylation143. In contrast, miRNAs in the cytoplasm exhibited an increased frequency of 3’ adenylation suggesting that terminal adenylation may antagonize terminal uridylation in this context.

Uridylation as a quality control mechanism

ncPAPs have well documented roles in the clearance of damaged nuclear ncRNAs in yeast144–

146. In humans, recent evidence stemming from studies focusing on the exosome-dependent

19 exonuclease DIS3L2 has uncovered a role for ZCCHC6/11-directed uridylation as a mechanism for clearance of damaged and dysfunctional ncRNAs in the cytoplasm. Identifying the RNA targets of exonucleases is technically challenging due the fact that these protein-RNA interactions result in the degradation of the RNA target. To overcome this obstacle, Pirouz et al. generated Dis3l2 knockout mouse embryonic stem cells and ectopically re-expressed a catalytic-dead mutant of

Dis3l2147. Through RNA-immunoprecipitation, they found that Dis3l2 binds numerous Pol III- transcribed ncRNAs including rRNAs, vault RNAs, and several classes of long-noncoding RNAs

(lncRNAs). Close analysis of the sequencing data revealed heavily uridylated 3’ ends and an enrichment for RNAs with mutations. Using similar strategies, two additional studies confirmed these observations in human cells and expanded the list of cytoplasmic ncRNAs targeted by this

TUTase/DIS3l2 axis to include Y-RNAs, tRNAs and a number of short Pol II-transcribed

RNAs148,149.

Uridylation-independent functions of the TUTases

Comparing the protein structure of ZCCHC6/11 to that of yeast homolog Cid-1 reveals that these human TUTases have acquired numerous additional domains over evolutionary time

(Figure 1.2). Notably, there are four zinc finger domains which have the potential to facilitate binding to a specific subset of RNA targets150 or to promote specific protein-protein interactions151,152. While the importance of domains outside of the catalytic site are not well understood, an N-terminal region of ZCCHC11 has reported functions that have been decoupled from its enzymatic activity153,154.

Human ZCCHC11 can attenuate the response to lipopolysaccharide stimulation in macrophages resulting in the decreased activation of the nuclear factor kappa-light-chain- enhancer of activated B cells (NF-kB)153. Surprisingly, while the catalytic domain was not required, overexpression of a portion of the N-terminus was sufficient to precipitate this phenotype.

Mechanistically, this likely occurs through protein-protein interactions between ZCCHC11 and

TIFA (TRAF-interacting protein with a forkhead-associated (FHA) domain), thus preventing TIFA- dependent activation of the kinase responsible for NF-kB activation. The N-terminus of ZCCHC11 was also shown to contribute to the regulation of the cell cycle in a number of human cell lines154.

Again, while the specific mechanistic details remain unclear, ZCCHC11 promotes the progression through the G1 phase of the cell cycle by enhancing the expression of cyclins and cyclin- dependent kinases, while ZCCHC11 knockdown results in slowed proliferation and accumulation of cells in the G1 phase.

1.5. Known biologic functions of TUTases.

Despite these insights into the biochemical consequences of TUTase-directed RNA uridylation, surprisingly little is known about the biological functions of these evolutionary conserved enzymes. Gain and loss-of-function studies have demonstrated that the TUTases can influence apoptosis through the global decay of mRNAs155, as well as the murine immune system likely through altered miRNA function72,156. Very recently, using transgenic murine lines in which both ZCCHC6 and ZCCHC11 were conditionally knocked out (DKO), it was shown that the

TUTases are required for oogenesis136. However, it remains to be seen if the control of mRNA turnover is a contributing mechanism in this context. Given that the regulation of mRNA decay has been shown to be an important regulatory mechanism in cellular differentiation and development157, it is tempting to speculate that the TUTases might therefore contribute to developmental processes beyond oogenesis. In support of this, morpholino-mediated depletion of both TUTases disrupted zebrafish embryonic development134. Similarly, the uridylation- selective exonuclease Dis3l2 was recently shown to be required for Drosophila gametogenesis158.

There is emerging evidence linking the TUTases and terminal uridylation to cancer. Depletion of ZCCHC11 has been shown to impair tumorigenicity and the metastatic potential of breast and ovarian cancer cell lines that express LIN28A through let-7-dependent mechanisms129. The let-7

21 family of miRNAs is an important set of tumor suppressors that acts by repressing oncogenes such as MYC, KRAS, and HMGA2159,160. The resulting increase in let-7 expression and the subsequent suppression of critical onocogenes is likely responsible for this effect. However, this study129 did not assess the contribution of ZCCHC6 either alone or in combination with ZCCHC11.

Given the redundancy of these TUTases and their contribution to regulation of miRNAs and mRNA stability, their role in cancer remains unclear. While numerous studies have linked disrupted mRNA stability to oncogenesis161, no report to date has directly addressed to potential contributions of uridylation-meditated mRNA decay in cancer.

In further support of a role for uridylation in cancer, germline loss-of-function mutations in

DIS3L2 result in Perlman’s Syndrome, a disease characterized by fetal overgrowth and increased susceptibility to Wilms’ tumor162. However, mechanistically it is unclear how loss of DIS3L2 results in these phenotypes. Finally, ZCCHC11 knockout mice exhibit impaired growth and disruption of insulin-like growth factor signaling132, a pathway with well-established roles in the regulating of cellular growth and metabolism in normal and oncogenic contexts163.

Despite recent insights into the contribution of terminal uridylation to gene regulation, the biological importance of ZCCHC6/11 in physiologic and pathologic settings remains poorly understood. To this end, this dissertation explores the role of these TUTases in normal development and in cancer.

AUTHOR CONTRIBUTIONS

This chapter is unpublished. D.S. Pearson wrote the text and generated the figures. A minor portion of Figure 1.4 was adapted from unpublished work from K.M. Tsanov. Edits were provided by P. Missios, T.E. North, and G.Q. Daley.

CHAPTER 2

The Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 Regulate Stem and

Progenitor Cell Fate in a Catalytic-independent Manner

2.1 Introduction.

Cellular differentiation is a complex process requiring the precise coordination of both intracellular and extracellular cues. The dramatic changes in gene expression driving embryonic development are often strikingly recapitulated during tissue regeneration in adults, as are the gene regulatory networks governing them164–167. While virtually every known layer of gene regulation from epigenetics168,169 to protein degradation170 have been implicated in the control of cellular differentiation, it is clear from studies of early mouse embryonic development171,172 as well as adult tissues stem cells173,174 that post-transcriptional control plays a prominent role.

Terminal RNA uridylation is a recently appreciated and poorly understood mechanism of gene regulation40,175. The enzymes responsible for carrying out uridine additions are a class of highly conserved non-canonical polyA polymerases (PAPs) called terminal uridylyltransferases

(TUTases)70,76. In contrast to canonical PAPs, which polyadenylate mRNAs co-transcriptionally in the nucleus, TUTases are thought to function primarily by adding non-templated uridines to the 3’ ends of a diversity of RNA substrates76,175,176. Emerging evidence suggests that TUTases regulate gene expression post-transcriptionally through their nucleotide transferase activity via a variety of mechanisms, including the clearance of aberrantly folded non-coding RNAs and the regulation of miRNA biogenesis and targeting40,147.

Among the most surprising, and recently discovered, targets of the TUTases are polyadenylated mRNAs100. In mammals mRNAs are primarily uridylated by two of the seven known TUTases, Zcchc6 and its paralog Zchcc11101. For simplicity, I will refer to Zchcc6 and

Zcchc11 collectively as the TUTases. While the biologic relevance of mRNA uridylation remains unclear, there is an association between uridylation frequency and decreased mRNA half- life100,101. Additionally, there is accumulating evidence in model organisms that mRNA uridylation is a widespread and evolutionary conserved mechanism of gene regulation with a range of molecular consequences (See Chapter 1)70,74,82,102. Additionally, ZCCHC11 has also been shown

24 to regulate cellular proliferation154 and the inflammatory response of macrophages153 independent of its uridylyltransferase activity, suggesting the potential for enzymatic-independent functions.

Zcchc6/11 are also known to function redundantly with the RNA-binding proteins Lin28a and

Lin28b in a uridylation-dependent manner to repress the biogenesis of the let-7 family of microRNAs (miRNAs)120,122,177. While both Lin28 and let-7 were originally discovered as regulators of developmental timing in C. elegans178, the Lin28/let-7 axis also contributes to the control of many processes including pluripotency120–123, tissue regeneration179 and metabolism180.

Additionally, Lin28a can promote skeletal muscle differentiation through increasing the translation efficiency of myogenic regulators including IGF-2181.

Gain and loss-of-function studies have demonstrated that the TUTases can influence apoptosis155, immune function72,153,156, oogenesis136, and zebra fish embryonic development134.

Additionally, a similar TUTase, Tailor, is required for Drosophila gametogenesis158. Collectively, this indicates that the TUTases might contribute to the regulation of mammalian development.

Consistent with this hypothesis, control of mRNA turnover has been shown to be an important regulatory mechanism contributing to cellular development and differentiation157.

Here, we report that Zcchc6 and Zcchc11 are highly expressed in many tissues during murine embryogenesis. TUTase protein expression is then dramatically downregulated across developmental time indicating a potential functional role in differentiation and development. To gain insight into this possibility, we explored the role of the TUTases in multiple in vitro models of cellular differentiation. We find that both Zcchc6 and Zcchc11 are highly expressed in lineage restricted muscle progenitors and that their expression is rapidly extinguished upon differentiation, similar to the changes observed in the developing embryo. Our functional characterization revealed that TUTases promote a less-differentiated cell state with ectopic expression of Zcchc6 impairing myotube formation and blunting the induction of differentiation-associated genes.

Surprisingly, overexpression of a catalytic-dead mutant of Zcchc6 also impaired differentiation,

25 implicating a role for uridylation-independent mechanisms in this process. Likewise, we find that overexpression of wild-type and catalytic dead TUTases only modestly impact mRNA half-lives, suggesting altered mRNA turnover is likely not responsible for this effect on differentiation. We then demonstrate that an N-terminal portion of Zcchc6 is required for the control of differentiation.

Finally, we show that depletion of TUTases in mouse embryonic stem cells (mESCs) promotes differentiation upon 2i/LIF withdrawal indicating that downregulation of the TUTases promote differentiation across diverse cell types of distinct lineages and developmental potencies. Taken together, these findings suggest that TUTases are likely broad regulators of cellular differentiation, in part, through uridylation-independent mechanisms of action.

2.2. Results.

TUTases are developmentally regulated genes

Zcchc6 and Zcchc11 are known to be expressed in mESC128. However, their expression pattern in later embryogenesis is unclear. To gain insight into the potential role of the TUTases in development, we profiled TUTase protein expression across five developmental time-points in seven different mouse tissues (heart, brain, liver, kidney, intestine, lung and stomach). Generally, both Zcchc6 and Zcchc11 proteins were highly expressed between embryonic days e12.5 and e14.5, followed by decreasing expression at e16.5 and e18.5 (Figure 2.1A). TUTase expression was comparatively lower or absent across all adult tissues, with the exception of persistent Zcchc6 expression in the liver. Western blot analysis of an additional 12 adult tissues revealed a similar pattern with >50% (11/19) expressing low or no expression of either TUTase (Figure 2.1C).

To shed light on the mechanism(s) by which the TUTases are regulated across development, we also assessed TUTase mRNA expression in these same tissues using qPCR. Strikingly, in our developmental time course, we found that expression of Zcchc6 mRNA did not correlate with protein expression across any tissue. In contrast, Zcchc11 mRNA tracked with

Figure 2.1. TUTases are developmentally regulated genes. (A) Western blot and (B) qPCR analysis of TUTase expression in embryonic and adult tissues isolated from CD1 mice. (C) Western blot and (D) qPCR analysis of Zcchc6/11 expression of additional adult tissues isolated from CD1 mice. Note, for Western blots, equal amounts of protein were loaded in each lane. For all qPCR experiments, relative expression was calculated by normalizing to the expression of either Gapdh or Actin corresponding to the protein loading controls shown in (A) and (C). For all adult time points, three independent mice were dissected. For all embryonic time points, tissues were pooled from multiple embryos to obtain sufficient material. qPCR samples represent RNA isolated from three independent mice or three independent pools of embryos. Error bars = standard error of the mean (s.e.m.).

27 protein expression in 4 of 7 tissues: brain, intestine, lung and stomach (Figure 2.6B). In agreement with previous reports of discordance between Zcchc11 mRNA and protein expression in adult mouse tissues,72 we conclude that TUTase expression is largely regulated at the protein level with the notable exception of skeletal muscle (Figure 2.6C &D). Consequently, TUTases appear to be developmentally regulated, primarily at the level of protein expression, in most tissues. We therefore hypothesized that TUTases play a functional role in regulation of development and cellular differentiation.

TUTases contribute to primary myoblast differentiation

Regeneration of mature skeletal muscle in response to injury has been widely studied as a paradigm for understanding the molecular mechanisms governing the intricate transitions from lineage-restricted stem cells to terminal differentiation. Additionally, the differentiation of adult satellite cells into mature muscle utilizes much of the gene regulatory mechanisms that control embryonic myogenesis164,182. We therefore decided to use in vitro models of muscle differentiation to test our hypothesis that the TUTases contribute to the regulation of cellular differentiation.

To determine if the TUTases play a functional role in muscle differentiation, we first profiled their expression in satellite-cell derived primary murine myoblasts (PM) and during their differentiation into mature myotubes (Figure 2.2A & B). Similar to what we observed in embryogenesis, proliferating PM exhibited robust expression of Zcchc6 and Zcchc11 and upon differentiation there was dramatic downregulation of both Zcchc6 and Zcchc11 protein expression

(Figure 2.2C). Additionally, we confirmed the induction of differentiation by blotting for two markers of mature myotubes: Creatine kinase (Ckm) and Myosin heavy chain (MHC). qPCR analysis further revealed a decrease of both TUTase mRNAs during differentiation suggesting that transcriptional/post-transcriptional mechanisms are responsible for this downregulation

(Figure 2.2D).

Figure 2.2. TUTases contribute to primary myoblast differentiation. (A) Schematic of myoblast differentiation into myotubes. (B) Culture conditions and corresponding bright-field images of primary myoblast differentiation. (C) Western blot analysis of Zcchc6/11, myosin heavy chain (MHC) and Creatine kinase (Ckm) across primary myoblast (PM) differentiation. (D) qPCR analysis of TUTase mRNA expression from (C). Relative expression was calculated by normalizing to the expression of Gapdh. (E) Western blot validation of PMs transduced with lentivirus containing empty-vector control (EV) or Zcchc6 cDNA (Z6OE) cDNA. (F) Bright-field images of EV and Z6OE on day minus one and day one of differentiation. (G) Immunofluorescence of EV and Z6OE cells stained for myosin heavy chain (MHC) or DAPI on day three of differentiation. (H) Quantification of MHC positive area from (G). (n =1). (I) Western blot analysis of Zcchc6/11 expression after transfection with control (NC) or Zcchc6/11 targeting siRNAs (DKD) on day one of differentiation. Cells were transfected two days prior to differentiation; double knockdown (DKD). (J) Bright-field images of NC and DKD cells on day minus one and day three of differentiation. (K) Immunofluorescence of NC and DKD cells stained for MHC or DAPI on day three of differentiation. (L) Quantification of MHC positive area from (K). (n =1). Note, scale bars = 250um, n.s = non-significant P>0.05; error bars = s.e.m.

Figure 2.2 (Continued)

Given a previous report of Lin28a-dependent regulation of myogenesis181, we additionally assessed Lin28a expression across PM differentiation. Consistent with this study, we detected very low expression of Lin28a mRNA on day four of differentiation by qPCR indicating that Lin28a is induced late in muscle differentiation (data not shown). However, Lin28a protein expression was below the detection limit by Western blotting across our entire differentiation time course

(Supplemental Figure 2.1A). The late induction of Lin28a mRNA, together with the early downregulation of TUTase expression, supports a predominantly Lin28a-independent role for the

TUTases in during muscle differentiation.

Given the downregulation of the TUTases observed during differentiation, we wondered if

TUTase overexpression would alter myogenesis. To test this, we stably overexpressed either

Zcchc6 or Zcchc11 in PMs by transduction with lentivirus. While we were able to successfully overexpress Zcchc6, no cells infected with viral particles for Zcchc11 survived the selection process, despite multiple independent attempts (Figure 2.2E and data not shown). Zcchc6 overexpression (Z6OE) impaired myotube formation relative to cells infected by empty vector control lentivirus (EV), which was evident as early as day 1 of differentiation (Figure 2.2F& G).

To quantify this effect, we utilized immunofluorescence to selectively stain myotubes with an antibody against the mature myotube marker myosin heavy chain (MHC)183. We then compared the total area of MHC positive staining and found that Zcchc6 overexpression reduced myotube formation (Figure 2.2F- H). Additionally, using siRNAs to knockdown both TUTases, we found that double knockdown (DKD) of Zcchc6/11 enhanced differentiation, indicating that the TUTases are regulators of myogenesis (Figure 2.2J-L).

TUTase regulation of myogenesis is uridylation-independent

To further investigate the role of the TUTases in myogenesis, we turned to C2C12 cells: an immortalized subclone184 of a mouse myoblast line originally derived from homogenized adult thigh muscle185. C2C12s are a commonly exploited in vitro system used to study the molecular

31 mechanisms of muscle differentiation due to their rapid proliferative capacity and ability to differentiate into mature myotubes, albeit at a much lower efficiency than PMs186. Additionally,

C2C12s do not express Lin28a181 and therefore can be used to further address the relationship between the TUTases and Lin28a. We profiled the expression of Zcchc6/11 protein expression by Western blotting across a time course of C2C12 differentiation. Similarly to PMs, Zcchc11 was downregulated upon myoblast differentiation, while Zcchc6 expression was unchanged (Figure

2.3A). The incomplete silencing of the TUTases during differentiation is not entirely surprising as it may reflect the relatively poor differentiation capacity of C2C12 cells relative to PMs186.

Myogenesis is composed of two temporally distinct processes: myoblast proliferation and differentiation. To determine the individual contribution of Zcchc6 and Zcchc11 to each stage, we knocked down the TUTases alone or in combination using siRNAs. In C2C12s, depletion of either or both TUTases had no impact on cellular proliferation suggesting that the TUTases are dispensable for myoblast proliferation (Supplemental Figure 2.1B). Because irreversible cell cycle exit occurs very early in myoblast differentiation187, unaltered cell proliferation upon knockdown reveals that TUTase depletion is not sufficient to induce differentiation. Next, we repeated the

TUTase knockdowns and induced differentiation 24 hours later. We found that depletion of either

Zcchc6 or Zcchc11 alone promoted, while double knock down (DKD) of Zcchc6/11 further enhanced, differentiation (Figures 2.3B & C). To rule out the possibility of off-target effects, we tested an additional two siRNAs targeting each Zcchc6 and Zcchc11. Consentient with our previous results, all siRNAs efficiently knocked-down their target and enhanced myotube formation to a similar degree (Supplemental Figure 2.1D & E). Taken together, these data indicate that the TUTases are dispensable for proliferating myoblasts, and that their downregulation promotes differentiation.

Next, we explored the impact of TUTase overexpression in C2C12s. Similar to PM, we were able to derive a line (Z6) with stable overexpression (around 3-4 fold above endogenous levels)

Figure 2.3. TUTase regulation of myogenesis is uridylation-independent. (A) Western blot analysis of Zcchc6/11 and myosin heavy chain (MHC) across C2C12 differentiation. Day zero indicates the switch from growth to differentiation media. (B) Immunofluorescence images of C2C12 cells transfected with the indicated siRNAs and stained for MHC or DAPI on day three of differentiation (n =3). (C) Quantification of MHC positive area from (H). (D) Western blot analysis of Zcchc6/11 expression after transfection with control or Zcchc6/11 targeting siRNAs on day four of differentiation. Cells were transfected two days prior to differentiation with either control (NC), Zcchc6 (Z6), Zcchc11 (Z11), or both (DKD) targeting siRNAs. (E) Western blot validation of C2C12 cells transduced with lentivirus containing empty-vector control (EV), Zcchc6 (Z6), or catalytic null Zcchc6 (Z6DD) cDNA. (F) Bright-field images of EV, Z6, and Z6DD C2C12s on day zero and three of differentiation. (G) Immunofluorescence of EV and Z6OE C2C12 cells stained for Myosin heavy chain (MHC) or DAPI on day four of differentiation. (H) Quantification of MHC positive area from (G). Note, scale bars = 250um; error bars = s.e.m.; **P < 0.01 and *** P < 0.001; n=3

33 of Zcchc6 (Figure 2.3E). Unfortunately, our attempts to overexpress Zcchc11 in C2C12s failed, for unclear reasons (data not shown). Similar to the siRNA-mediated TUTase depletion, Z6 cells proliferated at a similar rate to EV cells (Supplemental Figure 2.1C). However, upon induction of differentiation, Zcchc6 significantly impaired myotube formation relative to the EV control (Figure

2.3E-H). This indicates that Zcchc6 overexpression only impacts myoblasts in the presence of a differentiation signal.

While C2C12s do not express Lin28a181, the TUTases have been implicated in Lin28a- independent regulation of the differentiation-associated let-7 family of miRNAs77. Therefore, we wondered if TUTase-dependent regulation of let-7 could be contributing to this phenotype. To address this possibility, we assessed the impact of Zcchc6 overexpression on mature let-7 levels across C2C12 differentiation. Interestingly, we observed no difference in mature let-7 level upon

Zcchc6 overexpression at any point during differentiation (Supplemental Figure S2.1F).

Collectively, these loss- and gain-of-function experiments indicate that the TUTases are key regulators of muscle differentiation, which likely act independently of effects on Lin28a/let-7.

To determine if Zcchc6 influences differentiation through RNA uridylation, we additionally overexpressed a previously characterized catalytic-null mutant of Zcchc6 (Z6DD)188. This mutant was generated by replacing two aspartic acid residues (DD) in the active site with two alanines, thus disrupting catalytic triad and completely ablating uridylytransferase activity. Strikingly, Z6DD lines also displayed impaired myotube formation relative to EV. Furthermore, this inhibition was more robust than the Z6 line, despite the transgenes being expressed at similar levels (Figure

2.3E-H). This indicates the impact of Zcchc6 overexpression on muscle differentiation occurs independently of its uridylytransferase activity.

TUTase overexpression blunts the induction of differentiation genes

We next turned to RNA-sequencing as an unbiased approach to further characterize the molecular impact of wildtype and catalytic-null Zcchc6 overexpression on muscle differentiation.

Poly(A)-selected RNA-sequencing was performed in triplicate with both proliferating and differentiated (day 2) C2C12s. Consistent with our previous observation that perturbation of

TUTase expression did not impact the proliferation of C2C12s, very few genes were differentially expressed when comparing EV to Z6 or Z6DD under growth conditions (Supplemental Figure

S2.2A & B). Furthermore, principal component analysis (PCA) revealed very few differences between these lines (Figure 2.4A). In agreement with this, unsupervised hierarchical clustering

(UHC) failed to group these lines by genotype indicating that TUTase overexpression minimally impacts the transcriptome of proliferating myoblasts (Supplemental Figure 2.2C).

In stark contrast, PCA and UHC revealed large differences between EV and Z6 upon differentiation (Figure 2.4A & Supplemental Figure 2.2C). In agreement with a more pronounced impact on differentiation (Figure 2.2H), Z6DD was an outgroup in UHC, but clustered closer to EV than Z6 by PCA (Figure 2.4A & Supplemental Figure 2.2C). Furthermore, we found that the vast majority these differentially expressed genes in Z6 were also up-regulated (207 of 246; 84%) and down-regulated (181 of 197; 92%) in Z6DD (Figure 2.4B), indicating that Z6 and Z6DD are inhibiting differentiation through similar mechanisms. Initially, it was surprising that Z6DD had many more significantly changed genes relative to Z6. However, upon closer inspection, it became clear that the genes changed in Z6DD were also altered in Z6 just to a lesser degree

(Supplemental Figure S2.2D).

Given the high degree of overlap between differentially expressed genes comparing Z6 and

Z6DD, we surmised that the magnitude of changed targets, rather than their identity, might account for the more dramatic phenotype Z6DD. To further explore this, we defined a list of high- confidence “differentiation” (up-regulated) and “myoblast” (down-regulated) genes by identifying

Figure 2.4. TUTase overexpression blunts the induction of differentiation genes. (A) Principal components analysis of poly(A)-selected RNA-sequencing data generated from empty vector (EV; circles), Zcchc6 overexpression (Z6; triangles), and Zcchc6 catalytic-dead (Z6DD; squares) C2C12 lines under proliferative (Day minus one; red) and differentiation (Day two; blue) conditions. (B) Venn-diagrams of the overlap of significantly changed genes between Z6 and Z6DD compared to EV at day two of differentiation. (C) Heat map showing unsupervised hierarchal clustering of the top 164 differentially expressed genes. (D) qPCR analysis of mRNAs for Creatine kinase (Ckm), Myosin heavy chain 8 (Myo8), Myozenin 2 (Myoz2), and Phosphoglucomutase 5 (Pgm5) from (C). Relative expression was calculated by normalizing to the expression of Gapdh. Statistical significance was calculated relative to EV on Day four. (E) Western blot analysis of myotube markers Zcchc6, myosin heavy chain (MHC) and Ckm across differentiation. (F) Gene ontology analysis of genes significantly downregulated in Z6 or (G) Z6DD relative to EV. Note, error bars = s.e.m; .n.s. = non- significant P>0.05; *P < 0.05, n=3.

Figure 2.4 (Continued)

37 differentially expressed genes between proliferating differentiated (Day 2) EV cells. When we restricted our analysis to only consider genes with a fold-change of >50% and a P-value <0.05

(after correcting for multiple testing using the Benjamini-Hochberg procedure), we found 164 differentially expressed genes. Hierarchical clustering on gene names revealed that the majority of differentially expressed genes are “differentiation” genes (Figure 2.4C). Furthermore, the vast majority of “myoblast” genes were successfully downregulated in Z6 and Z6DD suggesting that

TUTase expression did not impair the silencing of the myoblast transcriptome.

Additionally, relative to EV, the induction of “differentiation” genes was blunted in Z6 and, to a greater degree, Z6DD (Supplemental Figure S2.2C). When considering the 150 most upregulated “differentiation” genes, 138 and 142 were blunted in Z6 and Z6DD respectively.

Furthermore, the degree of blunting was greater in Z6DD in all but 13 of these cases. When only considering the top 25 “differentiation” genes, all 25 followed this same pattern (Supplemental

Figure S2.2E).

To validate these results, we chose four of the top 25 differentially expressed genes (Creatine kinase (Ckm), Myosin heavy chain 8 (Myo8), Myozenin 2 (Myoz2), and Phosphoglucomutase 5

(Pgm5)) and assessed their expression by qPCR across an extended differentiation time-course.

In all four cases, Z6 impaired, while Z6DD further blunted, the induction of these “differentiation” genes (Figure 2.3D). Additionally, we confirmed the blunting Creatine kinase and Myosin heavy chain at the protein level by Western blot (Figure 2.4E).

In further support of this functional role, gene ontogeny (GO) analysis189,190 of significantly down-regulated genes between EV and Z6 or Z6DD showed enriched for many categories related to muscle differentiation and development (Figure 2.4F & G). GO analysis of upregulated genes revealed mostly unrelated categories such as “blood vessel morphogenesis” and “urogenital system development” (Supplemental Figure S2.2F & G). Curiously, there were also a number of enriched GO categories associated with osteogenesis including: “ossification” and

“odontogenesis”, perhaps relating to the capacity of mesodermal C2C12s to differentiate towards

38 an osteoblast lineage through using similar culture conditions191. Taken together, these data indicate that Zcchc6 overexpression blunts the induction of “differentiation” genes resulting in what is likely a delay of differentiation, while the silencing of “myoblast” genes is largely unaffected.

TUTase overexpression modestly impacts mRNA half-lives

The TUTases have been recently linked to the regulation of mRNA turnover101 and there is an emerging role for altered mRNA stability in the control of muscle differentiation35,192–194. Therefore, we explored the impact of Zcchc6 overexpression on mRNA half-life. Because mRNA half-lives change throughout differentiation35, we assessed the impact of Zcchc6 overexpression in proliferating myoblasts to prevent the differences in the differentiation state from confounding the half-life analysis.

We first treated proliferating EV, Z6 and Z6DD lines with actinomycin D to block transcription.

Then, we performed RNA-seq in triplicate on samples collected 0, 2 and 4 hours later and calculated mRNA half-lives following previously defined methods101 (Supplemental Figure S2.3A).

After stringent filtering, we were able to determine the half-lives of approximately 1,756 genes. To our surprise, only a handful of mRNAs had altered turnover in the Z6 line. However, for mRNAs with significantly altered half-lives, overexpression increased turnover with 27 of 45 and 40 of 54 mRNAs exhibiting reduced half-lives in Z6 and Z6DD respectively (Supplemental Figure S2.3B &

C). Additionally, the overlap of genes with significantly changed half-lives between Z6 and Z6DD was very poor (Supplemental Figure S2.3D & E). Collectively, these data indicate that TUTase overexpression only minimally impacts mRNA turnover under proliferating conditions. Thus, altered mRNA turnover is likely not responsible for Zcchc6’s impact on muscle differentiation.

An N-terminal region of Zcchc6 is required for the regulation of myogenesis

Zcchc6 is a large protein with multiple domains including four zinc fingers and a second catalytically inactive domain near the N-terminus (Figure 2.5A; See chapter 1 for extended description). To determine which region of the Zcchc6 protein is responsible for influencing myogenesis, we generated four truncation mutants of Z6DD (Figure 2.5A). We then derived

C2C12 lines which stably overexpress each mutant. Differentiation of these lines revealed that

Figure 2.5. The N-terminus of Zcchc6 is required for the regulation of myogenesis. (A) Schematic of truncation mutations for Zcchc6. Note, PAP (PolyA-polymerase). (B) Immunofluorescence images of the indicated C2C12 lines stained for MHC or DAPI at day three of differentiation. (C) Quantification of MHC positive area from (B) (n =2). Note, error bars = s.e.m. two Zcchc6 fragments (F1 & F4) inhibited myotube formation, while the other two fragments (F2

& F3) had no impact (Figure 2.5 B & C). F1 & F4 are terminal truncations of N- and C-termini indicating that these regions are dispensable. F2 further truncates the N-terminal portion of

Zcchc6 which includes a second predicted inactive catalytic domain indicating that this region is

40 required for the regulation of myogenesis. Consistent with this, F3 further truncates the N- terminus by deleting the intervening region between the two catalytic domains and also fails to suppress myotube formation. Thus, overexpression of Zcchc6 requires an N-terminal region for the inhibition of muscle differentiation.

TUTases regulate the exit from naive pluripotency

To determine if TUTase regulation is a more widespread phenomenon, we assessed their function in pluripotency and early differentiation using mESCs. mESCs grown in 2i/LIF conditions are in a stable “ground” state of pluripotency that molecularly resembles the pre-implantation epiblast195,196. Removal of these inhibitors results in rapid multi-lineage differentiation, which allowed us to interrogate the function of the TUTases in the transition out of pluripotency197,198.

We found that v6.5199 mESCs cultured in 2i/LIF highly express both TUTases and, similar to our observations during myogenesis, both Zcchc6/11 proteins are downregulated upon inhibitor removal (Figure 2.6A). We wondered if TUTase overexpression could impact the pluripotent state. To test this hypothesis, we generated doxycycline-inducible human ZCCHC6 and

ZCCHC11 overexpression (iZ6 and iZ11) mESCs lines using the KH2 system200. When cultured in more permissive conditions (mouse embryonic feeders and serum/LIF), induction of TUTase expression in iZ6 and iZ11 cells correlated with more intense alkaline phosphatase (AP) staining and more compact colony morphology, compared to un-induced controls (Figure 2.6B & C). This finding indicates that TUTase overexpression might be stabilizing the pluripotent state by transitioning cells closer to the “ground” state, despite the serum/LIF culture conditions. However, qPCR analysis of a panel of pluripotency markers upon ZCCHC6 or ZCCHC11 overexpression revealed virtually no differences (Supplemental Figure S2.4A & B). Furthermore, proliferation analysis showed that TUTase overexpression slowed growth, likely accounting for the smaller colony sizes observed upon doxycycline treatment (Supplemental Figure S2.4C). These

Figure 2.6. TUTases contribute to the exit from naïve pluripotency. (A) Western blot analysis of Zcchc6/11 expression in v6.5 mESCs upon removal of 2i/LIF. (B) Alkaline phosphatase staining of doxycycline-inducible ZCCHC6 (iZ6) and (C) ZCCHC11 (iZ11) overexpression mESC lines cultured in serum/LIF. Cells were treated with doxycycline (+dox) for 5 days prior to staining. Corresponding Western blot analysis of TUTase expression is shown below bright field images. (D) qPCR analysis for the pluripotency markers Nanog and (E) Tbx3 and (F) the differentiation marker Otx2 in iZ6 mESCs with or without dox upon 2i/LIF withdrawal (n=2). (G) Alkaline phosphatase staining and Western blot analysis of Zcchc6/11 from two independently derived mESC clones generated with non-targeting gRNAs (NTC) or Zcchc6/11-targeting gRNAs (DKO). Cells were cultured in serum/LIF conditions. (H) qPCR analysis from NTC and DKO cells for the pluripotency markers Oct4, (I) Nanog and (G) Rex1 upon 2i/LIF withdrawal (n=2). (K) Schematic of clonogenic 2i/LIF withdrawal assay. (L) Alkaline phosphatase (AP) staining of mESCs transfected with control (NC) or Zcchc6/11 targeting siRNAs (DKD) after 0 or 48hrs after withdrawal from 2i/LIF. (M) Quantification of AP positive colonies from (L). Note, error bars = s.e.m.; n.s = non-significant P>0.05; *P < 0.05, **P < 0.01 and *** P < 0.001. (n=3).

Figure 2.6 (Continued)

43 results indicate that overexpression of the TUTases is not sufficient to alter the pluripotent state under serum/LIF conditions.

To test the possibility that TUTase expression influences the exit from pluripotency, we differentiated iZ6 and iZ11 cells by removing 2i and LIF from the base media (N2B27). It is known that shortly after 2i/LIF removal the transcriptional program supporting pluripotency rapidly collapses with a concurrent induction of genes important for early differentiation198,201. We found that overexpression of ZCCHC6, but not ZCCHC11, impaired the reduction of the pluripotency factors Nanog and Tbx3 (Figure 2.6D & E and data not shown) and blunted the induction of the early commitment marker Otx2 (Figure 2.6F). This implies that ZCCHC6 is regulating the exit from pluripotency. Similarly, as we observed with myoblasts, when ZCCHC6 is overexpressed, a phenotype is revealed upon differentiation.

Next, we assessed the impact of TUTases loss on the maintenance of pluripotency under steady-state conditions. Using CRISPR-Cas9 and gRNAs targeting both TUTases, we were able to generate two independently-derived Zcchc6/11 double knockout (DKO) v6.5199 mESC lines

(Figure 2.6G). When maintained under permissive conditions (serum/LIF), both DKO lines had similar AP staining relative to controls (Figure 2.6G). This iindicats that the TUTases are dispensable for the maintenance of pluripotency.

To test the possibility that TUTase loss influences the exit from pluripotency, we differentiated

NTC and DKO cells by removing 2i and LIF. Consistent with our results from overexpression of

ZCCHC6 in the iZ6 line, we found that the pluripotency genes Oct4, Nanog, and Tbx3 decreased more rapidly in DKOs relative to NTCs. (Figure 2.6H-J). This further supports the conclusion that

TUTases regulate the exit from pluripotency.

To determine if the TUTases functionally influence the exit from pluripotency, we utilized a clonogenic commitment assay197. Briefly, 2i/LIF inhibitors are removed for 0, 16, 24 or 48hrs to

44 induce differentiation. Cells are then re-plated back into 2i/LIF conditions and the number of cells re-entering the pluripotent state are assessed by alkaline phosphatase staining of colonies arising

4-6 days later (Figure 2.6K). In a pilot study, we found significant clone-to-clone variability in the adherence of colonies arising after 2i/LIF withdrawal that did not correlate with TUTase expression status, thus making quantification unreliable (data not shown). To avoid this confounding variable, we instead used siRNAs to knockdown the TUTases. Depletion of the

TUTases decreased colony formation 48 hours after 2i/LIF withdrawal relative to cells transfected with control siRNAs (Figure 2.6L & M). This further supports the conclusion that TUTases regulate the exit from pluripotency. These data, together with our expression profiling in embryogenesis and our functional work in myogenesis, demonstrate that the TUTases are likely broad regulators of differentiation across diverse tissues and developmental stages.

2.3. Discussion.

Despite recent advances in uncovering the role of 3’ RNA uridylation in the regulation of gene expression, the biologic importance of the TUTases remains poorly understood. Here, we identify

Zcchc6 and Zcchc11 as developmentally regulated genes that influence the differentiation of pluripotent stem cells and muscle progenitors. Additionally, we show in both systems that

TUTases are dispensable for proliferating cells, thus highlighting their role as regulators of cell fate transitions.

In myoblast differentiation, Zcchc6 overexpression blunted the induction of “differentiation” genes. Interestingly, catalytic activity was dispensable for this effect. While we demonstrate that the N-terminus of Zcchc6 is required for this effect, the mechanism(s) by which this region impacts differentiation remains unclear. It would be insightful to further truncate Zcchc6 from the C- terminal end to determine a minimal region that is sufficient to impair myogenesis. Similarly, additional N-terminal truncations will be required to define the minimal portion of N-terminus that

45 is required to inhibit myogenesis. However, this is not the first report of an enzymatic-independent function of a TUTase. In the mouse macrophage cell line RAW264.7, ectopic expression of an

N-terminus portion of human ZCCHC11 was sufficient to attenuate the response to lipopolysaccharide stimulation through decreased activation of NF-kB153. Mechanistically, this likely occurs through protein-protein interactions between ZCCHC11 and TIFA (TRAF-interacting protein with a forkhead-associated (FHA) domain), thus preventing TIFA-dependent activation of

IkB kinase (IKK) the enzyme directly responsible for NF-kB activation. Similar to our observations, the catalytic domain of ZCCHC11 was not required, while a portion of the N-terminus was sufficient to precipitate this phenotype. Additionally, the N-terminus of ZCCHC11 was also shown to be required for the regulation of cell cycle in different human cell lines154. Again, it is unclear how the N-terminal domain carries out these functions. Therefore, it is of interest to define the protein-protein interactions of the N-terminus of Zcchc6. Given the lack of a proliferative phenotype upon either TUTase overexpression or knockdown, it would be informative to perform these experiments in both proliferating and differentiating myoblasts.

Multiple lines of evidence indicate that TUTase function in muscle differentiation is independent of Lin28a/let-7 axis. First, Lin28a and the TUTases are never expressed at the same time in either system. In PMs, TUTases are highly expressed under proliferating conditions and their expression is rapidly extinguished upon differentiation. In contrast, Lin28a expression has been reported to be induced late in muscle differentiation in PMs, while in C2C12s differentiation does not induce Lin28a expression181. Consistent with this, we were only able to detect Lin28a expression in PMs by qPCR in the last time point of differentiation (data not shown). Second, ectopic expression of Lin28a and the TUTases have opposite phenotypic effects. Lin28a overexpression promotes, whereas TUTase overexpression inhibits, differentiation. Finally, we show that TUTase expression has no impact on mature let-7 expression in either proliferating or

46 differentiating C2C12s. Taken together, it is clear that the TUTases regulate muscle differentiation independent of Lin28a.

In mESCs, TUTase depletion similarly promotes differentiation. However, the mechanistic relationship between the TUTases and Lin28a is less clear. mESCs express Lin28a under 2i/LIF conditions and it has been suggested that TUTases contribute to the regulation of pluripotency through let-7 in this context202. It would be interesting to repeat TUTase loss-of-function experiments using Lin28a/b DKO lines to clarify the nature of this relationship203. Our results are consistent with a recent report in which the TUTases were reported to be dispensable for mESCs cultured under serum/LIF conditions136. While this study did not assess the kinetics of differentiation, they did show that TUTases DKO mESCs were capable of forming all three germ layers. Taken together with the data presented here, the TUTases can enhance differentiation and the tissues that arise from DKO mESCs are likely identical to those from control lines.

While enhanced differentiation due to TUTase loss might be well tolerated in vitro, embryonic development is the result of countless and carefully coordinated cell fate transitions that require the integration of many cell autonomous and non-autonomous signals. Therefore, the TUTases might contribute to embryogenesis by ensuring that the timing of critical cell fate transitions occurs with high precision. Single knockout lines for Zcchc6 and Zcchc11 are viable, yet exhibit defects in the immune system156 and disrupted insulin signaling respectively132, indicating that single

TUTase knockout is tolerated during embryogenesis. However, given the redundancy of TUTase function reported here and elsewhere101,128, elucidating the function of TUTases in embryonic development remains of great interest, but will likely require the use of tissue specific and/or developmentally restricted cre recombinase activity.

Finally, given the broad developmental expression pattern of the TUTases and the molecular similarities of many developmental and regenerative programs164–167, it would be interesting to interrogate the role of the TUTases in models of tissue regeneration. Our results implicating the

TUTases in the regulation of myogenesis also suggest that their inhibition might enhance skeletal muscle regeneration in response to injury in vivo. Furthermore, TUTase inhibition maybe useful in those contexts to generally enhance muscle differentiation. However, instances where impaired differentiation of myoblasts has been implicated in disease etiology, such as myotonic dystrophy204, should be prioritized. More generally, it would be fascinating to see if TUTases contribute to the maintenance of other tissues through the regulation of the differentiation of tissue-restricted stem cells.

2.4 Methods.

Plasmids

Stable overexpression of Zcchc6 was achieved by amplifying FLAG-tagged cDNAs from pcDNA3-

FLAG-mTUT7 (a gift from Z. Mourelatos; Addgene plasmid #60044) then cloning the purified amplicon into pLentiCas9-Blast using the In-Fusion HD cloning system (Clontech) after excision of Cas9 by digestion with NheI and BamHI. To generate an empty vector control, pLentiCas9-

Blast was digested with NheI and BamHI, blunted with a Klenow enzyme (NEB), and re-ligated.

Mutations to the catalytic domain of Zcchc6 were introduced using the Quikchange ii mutagenesis kit (Agilent) with the following primers: F: 5'-

CATGTCCGTTAATCGTCATACAGACAGCAAGAGCACTCTGTTTGAACCCAAATCCATT-3'; R:

5'-AATGGATTTGGGTTCAAACAGAGTGCTCTTGCTGTCTGTATGACGATTAACGGACATG-3'

Doxycycline-Inducible mESC lines expressing either human ZCCHC6 or ZCCHC11 cDNAs were generated using the KH2 system200. Briefly, FLAG-tagged TUTase cDNAs were amplified from cDNA libraries generated from oligo-dT primed reverse transcription (Super Script III; Invitrogen) using RNA isolated from Hct116 cells as a template. Amplicons were then cloned into the pBS31 plasmid (Open Biosystems #MES4486) and purified using isopropanol precipitation. Purified plasmids were delivered to parental KH2 cells, along with pCAGGS-FLPe (Open Biosystems

#MES4488), via electroporation (Lonza #vaph-1001). To generate clonal TUTase KO mESC lines, we utilized gRNAs against mouse Zcchc6, Zcchc11 and two non-targeting controls

(NTC)205. One NTC and mouse Zcchc6 gRNA was cloned into pSpCas9-2A-GFP while the other

NTC and Zcchc11 gRNAs were cloned into pSpCas9-2A-mCherry. The sequences of the gRNAs are as follows: (NTC1: GTAGGCGCGCCGCTCTCTAC and NTC2:

GGGCCCGCATAGGATATCGC). mZcchc6-1: GTGCTTATGAGCAAACGGAA mZcchc11-1:

AATCCGCCAGGACATTGTGG. gRNAs were selected using the CRISPR Design Tool according to the website’s recommendations (http://crispr.mit.edu/). The lentiviral packaging plasmids pMD2.G (Addgene plasmid #12259) and psPAX2 (Addgene plasmid #12259) were gifts from D.

Trono. pLentiCas9-Blast (Addgene plasmid #52962) and pSpCas9-2A-GFP (Addgene plasmid

#48138) were gifts from F. Zhang and pSpCas9-2A-mCherry was a gift from and M. Stitzel.

Transfections

1000ng of each pSpCas9(BB)-2A-GFP and pSpCas9(BB)-2A-mCherry were transfected using

Lipofectamine 2000 according to the manufacturer’s protocol. siRNA transfections into a well of a 6 well plate were conducted as follows: 1.5 x 105 primary myoblasts, V6.5 or C2C12 cells were reverse transfected with 80 pmol, 80 pmol and 40 pmol of each siRNA respectively using 5ul of

Lipofectamine RNAiMAX (Invitrogen). Transfections were scaled to fit other well sizes but keeping the ratios of siRNA:RNAiMAX:Cell number constant. In double-knockdowns experiments, the control siRNA volume was doubled to correspond the total siRNA volume in the knockdown condition. The following siRNAs were used (Note that unless otherwise specified mZ6-1 and mZ11-2 were used):

SiRNA Name Catalog Number NC Ambion #4390843 mZ6-1 Ambion #4390815-s103051 mZ6-2 Ambion #4390815-s103052 mZ6-3 Ambion #4390815-s103053 mZ11-1 Ambion #4390815-s106496

mZ11-2 Ambion #4390815-s106497 mZ11-3 Ambion #4390815-s106498

Cell Culture

Satellite cell-derived myoblasts were a gift from A.J. Wagers that were isolated as previously described206 and maintained in 20% F10 media supplemented with 20% horse serum, 1% glutaMAX (Thermo Scientific A1286001), 1 U/ml Penicillin and 1 μg/ml Streptomycin on plates coated with a mixture of 1 ug/mL Collagen Type I from Rat Tail (Sigma C7661), 10 ug/mL mouse

Laminin (Invitrogen #23017-0155). Growth media was changed every other day and 5 ng/mL bFGF (Peprotech #100-18B) was added daily. Cells were passaged every 2-5 days using 5mM

EDTA/PBS (Thermo Scientific AM9260G). C2C12 (ATCC-CRL-1772) cells were grown in DMEM supplemented with 10% FBS, 2mM glutamate, 1 U/ml Penicillin and 1 μg/ml Streptomycin. For differentiation of both primary myoblasts and C2C12, growth medium was replaced with DMEM with 2% horse serum (Thermo Scientific #26050088), 1% glutaMAX, 1 U/ml Penicillin and 1 μg/ml

Streptomycin. v6.5199, KH2200 doxycycline-inducible iZ6 and iZ11, and NTC and TUTase DKO mESCs were grown on irradiated CF1 MEFs (GlobalStem) in mESC medium (DMEM, 15% FBS,

1 U/ml Penicillin, 1 μg/ml Streptomycin, 2 mM L-glutamine, 0.1 mM NEAA, 0.1 mM BME, 1,000

U/ml LIF) or in 2i/LIF media195 on laminin coated plates (Sigma # L2020). iZ6 and iZ11 cells were generated using KH2 cells as described above and inducible expression was validated by treatment with 2 ug/ml doxycycline for 72hrs followed by Western Blotting. NTC and DKO cells were derived from parental v6.5 cells by FACS for double positive cells 24-72 hrs after co- transfection of pSpCas9-2A-GFP and pSpCas9-2A-mCherry. For mESCs plated on MEFs in mESC medium, cells were grown for five days, and stained using the Leukocyte Alkaline

Phosphatase kit (Sigma # 86R) following the manufacturer’s instructions. In 2i/LIF withdrawal experiments, 2i and LIF were excluded from the medium 24 hrs post-siRNA transfection. At the

50 specified intervals 250 cells were plated back into 12-well plates with 2i/LIF medium, grown for 4-

6 days and then stained for alkaline phosphatase.

Stable Cell Line Generation

Stable cell lines were generated by infection with un-concentrated viral supernatant and selected with either 1 ug/ml puromycin or 7.5 ug/ml blasticidin. Briefly, viral supernatant was centrifuged at

1,200 RPM for 5 min to pellet cell debris and used immediately or stored at 4°C for later use. To produce lentiviral supernatant, 2.5 x 106 293T cells were plated in a 10-cm plate and transfected the next day using Lipofectamine 2000 and OptiMEM (Life Technologies). For a 10cm plate, a mixture containing 6.6 ug transfer plasmid, 3.3 ug pMD2.G, 4.8 ug psPAX2, 36 ul Lipofectamine

2000, and 600 ul OptiMEM was used. After 12-16 hrs, medium was changed to DMEM/20% FBS.

Viral supernatant was collected at 24 and 48 hrs.

Cell Proliferation Analysis

Rates of cellular proliferation were determined by staining with trypan-blue and counting live cells with a hemocytometer (Thermo Fisher Scientific # 22600100). C2C12 cells were seeded 5.0 x

105 cells/well on two 12-well plates. Day 0 counts were obtained 12-24hrs later to ensure equal starting numbers. The second plate was counted 96 hrs after Day 0 with cells being split as need to prevent reaching confluency. Proliferative rates were calculated by multiplying 96hr counts by split ratios and then by dividing by Day 0 counts. Final proliferative values were calculated by normalizing to the appropriate controls.

Immunofluorescence

Satellite cell-derived myoblasts were plated at a density of 4,000 cells/well (C2C12 at 3,000 cells/well) in 96-well imaging microplates (Corning #353219) and differentiated as described above. In the case of siRNA knockdown experiments, cells were reverse transfected and plated

51 directly into 96-well plates (Falcon #353219). Cells were then fixed for 15 minutes at room temperature in 4% paraformaldehyde diluted in PBS and then blocked in 0.1% triton x-100, 1%

BSA for 30 minutes. Cells were probed with anti-myosin antibody (MF-20,DSHB ;1:250) overnight at 4°C and then stained with Alexa Fluor 568 (Life Technologies; 1:1000) for 1hr at room temperature followed by co-staining with DAPI.

Myotube Image Acquisition/Analysis

Differentiated myoblasts were imaged on Cell Voyager 7000 (Yokogawa). Five images per well were taken at 10X magnification using each fluorescent channel. Images were processed using

FIJI207 as previously described with minor modifications183. Briefly, threshold intensities were set manually for each plate using three randomly selected wells. To eliminate spurious background, particles smaller than 2.0 um2 were discarded. Total area positive MHC area was then calculated for each field-of-view and then averaged per well. For each condition at least five wells were imaged and for each data set, unless otherwise noted, at least 3 biologic replicates were analyzed.

In PM experiments adhesion of mature myotubes was poor. Analysis was performed only on images where there was minimal evidence of myotube detachment.

Mouse Tissue Isolation

Embryonic tissues were isolated from embryos produced from timed matings of CD-1 mice

(Charles River). Specific tissues were identified by morphology and excised under a dissection microscope. Adult mouse tissues harvested in a similar manner using 6-10 week old CD-1 mice.

All animal experiments were in accordance with guidelines approved by Boston Children’s

Hospital Institutional Animal Care and Use Committee.

Western Blotting

All tissues and cells were lysed on ice in RIPA buffer supplemented with HALT-protease inhibitors

(Pierce). Primary tissues were additionally subjected to sonication using a Bioruptor (Diagenode).

Proteins were denatured using DTT and LDS according to the manufacturer’s instructions

(Thermo Scientific NP0004) then run on a 4-20% polyacrylamide gel (Bio-Rad) and transferred to a PVDF membrane (Millipore). Membranes were blocked for 60 minutes in either 5%milk/PBST

(chemiluminescent blots) or 3%BSA/PBS (fluorescent blots). Primary antibodies resuspended in

3%BSA/PBS were incubated with membranes at 4°C overnight on a shaker. Secondary antibodies were incubated for 1-4 hours at room temperature or 4°C overnight. Protein expression was assessed by chemiluminescence using SuperSignal West Pico and Femto reagents (Thermo

Scientific) or by fluorescence using Odyssey CLx imaging system (LI-COR). The following antibodies were used:

The following secondary antibodies used were used for chemiluminescence: HRP-anti-rabbit IgG

(GE Healthcare #NA934, 1:2,000), anti-mouse IgG (GE Healthcare #NA931; 1:2,000), or anti- goat IgG (Santa Cruz sc-2020; 1:2,000). Secondary fluorescence antibodies: IRDye-680RD anti- rabbit IgG (LI-COR #925-68071; 1:20,000), IRDye-680RD anti-goat IgG (LI-COR #925-68074;

1:20,000) and IRDye-800CW anti-mouse IgG (LI-COR #925-32210; 1:20,000). Image Studio was used to quantify fluorescent blots following the manufacturer’s guidelines (Odyssey CLx (LI-

COR)).

Quantitative PCR (qRT-PCR)

Total RNA was isolated using Trizol (Invitrogen) combined with miRNeasy columns (Qiagen) according to the manufacture’s protocols. Reverse-transcribed was performed on 100-500ng total

RNA using the miScript II RT kit using the hiFlex buffer system (Qiagen). cDNAs were subjected to SYBR green-qPCR using in house qPCR primers or pre-designed primer assays for U6 snRNA or miScript miRNA assays (Qiagen) (miRNAs). RNA expression was calculated using the ΔΔCt method using either Gapdh or β-actin for normalization. qPCR primer sequences were designed using the IDT PrimerQuest design tool (IDT, USA) or were obtained from Zeisel et al.’s qPCR primer database208. The following primers were used in this chapter:

Primer Name Sequence mGapdh-F CATGGCCTTCCGTGTTCCT mGapdh-R GCGGCACGTCAGATCCA mActb-F CAGAAGGAGATTACTGCTCTGGCT mActb-R TACTCCTGCTTGCTGATCCACATC mZcchc6-F CAGTCAGGTAGCCTTTCCAGTA mZcchc6-R GCAGTTCCTTCCCTCATGATTC mZcchc11-F TCTATGCTCAAGCAGACAGATG mZcchc11-R ACTGACACTGAGGTACGGATA mCkm-F AGACAAGCATAAGACCGACCT mCkm-R AGGCAGAGTGTAACCCTTGAT mMyh8-1F GGAGGACCAAATATGAGACCG mMyh8-1R GTTCACGGCTTCTACGTGCT mMyoZ2-1F CCCAAATCCAGAGAACATCG mMyoZ2-1R TCCCATGGAGACCGGTAGTA mPgm5-1F CGATGCTCTCTGTGTGGAGA mPgm5-1R AATGGACAGCCAGACCAAGA mOct4-F TGGATCCTCGAACCTGGCTA mOct4-R CCCTCCGCAGAACTCGTATG mNanog-F AATGCTGCTCCGCTCCATAA mNanog-R TAAAATGCGCATGGCTTTCC mRex1-F GGCCTCTTTTGGTATTCCATGG mRex1-R CCCATCCCCTTCAATAGCACAT mKlf2-F TAAAGGCGCATCTGCGTACA mKlf2-R CGCACAAGTGGCACTGAAAG mKlf4-F GCACACCTGCGAACTCACAC mKlf4-R GTTTGCGGTAGTGCCTGGTC mKlf5-F TTGCTTCCAAACTGGCGATT mKlf5-R AGGTGGGAGAGTTGGCGAAT mTbx3-F AGCGGGGTACAGAGATGGTC

mTbx3-R TTGGCCTTTTTATCCAGTCCAG mTfcp2l1-F CAGCCCGAACACTACAACCAG mTfcp2l1-R CAGCCGGATTTCATACGACTG mOtx2-F TATCTAAAGCAACCGCCTTACG mOtx2-R GCCCTAGTAAATGTCGTCCTCTC mOct6-F TACCGCGAAGTGCAGAAGC mOct6-R CGTGGGTAGCCATTGAGGG mLin28a-F AGGCGGTGGAGTTCACCTTTAAGA mLin28a-R AGCTTGCATTCCTTGGCATGATGG mLin28b-F GCATGGGATTCGGATTCATCT mLin28b-R CCTTCCATGAATAGTTTGCTTTGG U6 Qiagen #MS00033740 Let-7a Qiagen #MI0000556 Let-7b Qiagen # MI0000558 Let-7c Qiagen # MI0000559 Let-7e Qiagen # MI0000561 Let-7f Qiagen # MI0000562 Let-7g Qiagen # MI0000137

RNA Sequencing

Total RNA (100-500ng) of RNA was enriched with either poly(A) selection (NEBNext Poly(A) mRNA Magnetic Isolation Module) or rRNA depletion (NEBNext rRNA Depletion Kit

(Human/Mouse/Rat). Libraries were constructed using NEBNext Ultra RNA Library Prep Kit.

Small RNA libraries were constructed using NEBNext Small RNA Library Prep Kit for Illumina.

Average library size was estimated using either a Bioanalyzer (Agilent) or 4200 Tapestation

(Agilent). Library concentration was determined using both qRT-PCR (Kapa Biosystems) and

Qubit dsDNA high-sensitivity assay (Invitrogen). Sequencing was performed on either the

NextSeq 500 or HiSeq 2500 platforms (Illumina) using 75-bp single-end reads. Gene expression

(RPKM) was determined with TopHat209 and HTSeq-count210. The DAVID 6.8 Functional

Annotation Tool was use for GO analysis189,190.

RNA half-life analysis

EV, Z6 and Z6DD C2C12 lines were plated in 6-well plates to achieve 40-60% confluence. Cells were treated with actinomycin-D (Thermo Scientific) and harvested as indicated in the text. RNA- library were constructed and mRNA half-lives were calculated all as previously described101.

Statistics

Statistical significance of P<0.05, P<0.01 or P<0.001 are denoted with one, two or three asterisks respectively. P values were calculated using a two-tailed Student’s t-test (unless otherwise specified). Data are shown as mean of biological replicates with error bars displaying variation as calculated by s.e.m. unless otherwise stated. In the case of RNA half-life measurements, paired t-tests were used and the Benjamini-Hochberg correction was applied to account for multiple hypotheses testing.

Data Availability

All data contained within this manuscript are available upon request. RNA-sequencing data sets will be deposited in the Gene Expression Omnibus (GEO) prior to publication.

AUTHOR CONTRIBUTIONS

This work is an iteration of a manuscript in preparation written by D.S. Pearson in close collaboration with K.M. Tsanov. D.S. Pearson and K.M. Tsanov designed the experiments. D.S.

Pearson wrote the text and generated most of the data. K.M. Tsanov contributed to Figure 2.3A,

Figure 2.6B-J and Supplemental Figure S2.4, aspects of the Methods section and a number of related pilot experiments not included in this work. M. Morris helped prepare the RNA-seq libraries (Figure 2.4) A. Han analyzed the RNA-seq data sets. Y.C. Huang helped with WB expression analysis, cell culture, and IF related to Figures 2.1, 2.2 & 2.5.. J.K. Osborne, P. Sousa,

56 and Y.C Huang assisted with the mouse tissue isolation. A. Han, J.T. Powers, P. Missios and

J.K. Osborne assisted with experimental design. G.Q. Daley designed and supervised experiments.

ACKNOWLEDGMENTS

We would like to thank: S. Buratowski, J. Rinn, H. R. Horvitz, and numerous members of the

Daley lab for many insightful discussions; T. Schlaeger (BCH) for technical assistance in IF image acquisition and analysis; R. Rubio CCCB (DFCI) and X. Wu/Yi Zhang laboratory for technical help in performing the RNA-seq; R. Mathieu and M. Paktinat for helping with flow cytometry. The

IDDRC Molecular Genetics Core (BCH) performed the Bioanalyzer analysis and was supported by NIH (NIH-P30-HD 18655). Intensive informatics analysis was performed utilizing the Orchestra

High Performance Computing Cluster (HMS). D.S. Pearson is supported by a grant from NIGMS

(T32GM007753). K.M. Tsanov was a Herchel Smith graduate fellow and a Howard Hughes

Medical Institute international student research fellow. G.Q. Daley was supported by the Howard

Hughes Medical Institute and a grant from the NIGMS (R01GM107536). G.Q. Daley also is an investigator of the Manton Center for Orphan Disease Research.

CHAPTER 3

The Role of the Terminal RNA Uridylyltransferases ZCCHC6 and ZCCHC11 in Cancer

3.1 Introduction.

In 2000, Hanahan and Weinberg proposed the existence of six hallmarks of cancer that are acquired over time and collectively account for the insidious nature of the disease211. From sustained proliferative signaling to the activation of invasive and metastatic properties, it is known that these features are acquired by normal cells primarily through genome instability212. The mechanisms by which genomic instability gives rise to these hallmarks has thus been the focus of intense study over the last two decades. It is now understood that dysregulation of transcriptional control is an inescapable consequence of genome instability and is directly involved in oncogenesis14. However, despite a growing appreciation for the role of mRNA turnover as a potent post-transcriptional mechanism of gene regulation157, the contribution of dysregulated mRNA decay in cancer is much less clear161,213,214.

ZCCHC6 and ZCCHC11 are terminal RNA uridylyltransferases that have been recently identified as regulators of mRNA turnover through their enzymatic activity101. Additionally, these particular TUTases have been implicated in cancer in the context of the LIN28/let-7 axis, where they act redundantly to reduce the expression of the tumor suppressor miRNA let-7 in a uridylation-dependent manner122,124,129,177. Depletion of ZCCHC11 has been shown to impair tumorigenicity and the metastatic potential of breast and ovarian cancer cell lines that express

LIN28A through let-7-dependent mechanisms129. However, the role of the TUTases in cancer outside the LIN28/let-7 axis, if any, has not yet been addressed.

For decades, it has been known that embryonic and developmental genes are frequently re- expressed in cancer215. This class of genes, termed “oncofetal”, are defined by their expression pattern in which there is expression during fetal development, largely no expression in the adult, and then reactivation of expression in cancer216. Recently, it has become clear that many oncofetal genes, such as c-MYC and Oct4, play critical functional roles in both development and oncogenesis217,218. In light of our recent work identifying the role of TUTases in the regulation of

59 normal cellular differentiation (Chapter 2), we wondered if TUTases might also be important regulators in cancer.

Here, we report that TUTases exhibit an oncofetal pattern of expression, in which they are robustly expressed across a diversity of human cancer cell lines and primary colon cancer tumor samples despite low expression in normal adult human tissues. Functionally, TUTase expression contributed to the growth and tumorigenicity of a subset of cancer cell lines, likely through

LIN28A/B and miRNA-independent mechanisms. Depletion of TUTases also resulted in altered mRNA uridylation and turnover, including the dysregulation of cell cycle factors and core histone proteins. Finally, we show that TUTase loss sensitized cancer cells to DNA-damaging agents, possibly due to disrupted histone turnover. Taken together, our results suggest an oncofetal function for the TUTases, and implicate TUTase-dependent mRNA turnover as a previously unappreciated vulnerability of cancer.

3.2. Results.

TUTases exhibit an oncofetal pattern of expression To determine the expression pattern of ZCCHC6/11 in human cancers, we first assessed protein expression by Western blotting across a panel of 14 normal adult tissues and 29 cancer cell lines representing 10 diverse types of cancer. Similar to that of the adult mouse (Figure 2.1C),

9 of 14 normal adult tissues expressed little or no ZCCHC6 (Figure 3.1A). ZCCHC11 exhibited a similar pattern to ZCCHC6, but was expressed in more than half of adult tissues (9 of 14). In contrast, ZCCHC6 and ZCCHC11 were highly expressed in nearly all cancer cell lines relative to normal fibroblasts (Figure 3.1B). When comparing normal adult tissues with low TUTase expression to corresponding cancer cell lines, such as those arising from intestine, colon and liver, we observed abnormal ZCCHC6 and ZCCHC11 expression in the cancer context. Further, in cancer cell lines, there was no correlation between TUTase and LIN28A/B expression indicating

Figure 3.1. TUTases exhibit an oncofetal pattern of expression. (A) Western blot analysis of TUTase expression in normal adult human tissues. (B) Western blot analysis of ZCCHC6/11 and LIN28A/B protein expression in human cancer cell lines. (C) RNA expression data for ZCCHC6/11 across 37 cancer types from TGCA visualized with FIREBROWSE. (D) Western blot analysis of ZCCHC6/11 from human colorectal tumors (T) and normal tissues (N) of unmatched samples (left) and matched tumor and normal adjacent tissue (right). Note, for Western blots equal protein was loaded per lane; FIREBROWSE: http://firebrowse.org/, RSEM (RNA-seq by Expected Maximization). Green arrows highlight colorectal cancer (CHOL). The identities of cancer cell lines in (B) are listed in the Methods section.

Figure 3.1 (Continued)

62 functional roles of the TUTases beyond the LIN28/let-7 axis (Figure 3.1B). We therefore conclude that the TUTases, in particular ZCCHC6, exhibit a classical oncofetal pattern of expression in some cancers.

To better understand the expression pattern of TUTases in cancer, we turned to publicly available RNA-sequencing data sets from The Cancer Genome Atlas (TGCA). Surprisingly,

ZCCHC6/11 mRNA expression in 37 human primary tumor types were quite similar to normal tissues of origin (Figure 3.1C). However, when considering the discordance we previously observed between TUTase RNA and protein expression across mouse development (Figure 2.1C

& D), we decided to further assess protein expression to more clearly determine instances of

TUTase reactivation. Therefore, we analyzed TUTase protein expression in a set of primary human colorectal tumors. Despite having no evidence for TUTase overexpression at the mRNA level in TGCA data sets (Figure 3.1C), 50% (5/10) of colorectal tumors exhibited ZCCHC6 and

ZCCHC11 overexpression at the protein level, compared to normal mucosa samples, which exhibited very low levels of TUTase protein expression (Figure 3.1D). Next, we analyzed data from 170 cancer studies available from TGCA to determine the mutational landscape of

ZCCHC6/11. Here, we found that ZCCHC6/11 do not harbor frequently recurring mutations, amplifications or deletions (Supplemental Figure S3.1A-C). Together, these data suggest that

ZCCHC6 and ZCCHC11 are overexpressed at the protein level in a wide-range of cancer cell lines and tumor types.

ZCCHC6/11 are dispensable for NIH3T3 transformation by KRASG12V

To explore the possibility that ZCCHC6/11 are functionally relevant in cancer, we first utilized

NIH3T3 cells as an in vitro system to assess the function of the TUTases in oncogenic transformation. NIH3T3 cells are immortalized fibroblasts that can be transformed by the

63 overexpression of a single oncogene such as mutant Kirsten Rat Sarcoma Viral Oncogene

Homolog (KRAS)219 or Erb-B2 Receptor Tyrosine Kinase 2 (erbB-2)220. We first generated clonal

ZCCHC6/11 double knockout (DKO) and control lines by transiently expressing Cas9 and

ZCCHC6/11-targeting and non-targeting control gRNAs (NTC). Utilizing Western blotting as a screen for successful ZCCHC6/11 knockout, we were able to generate three independent DKO lines (Supplemental Figure S3.2A). Consistent with our previous observations in normal mouse embryonic stem cells and muscle progenitors, we found no significant difference in the proliferative rates of NIH3T3 NTC and DKO lines (Supplemental Figure S3.2B).

To directly assess the role of TUTases in oncogenesis, we transformed NTC and DKO lines using retrovirus transduction to stably overexpress a constitutively active mutant of KRAS

(KRASG12V)221 and then assessed growth in vivo using allograft assays. For this study, we injected transformed NTC and DKO cells subcutaneously into the flanks of immunocompromised (Rag2-/- gc-/-) mice and allowed them to grow for 17 days. We found that DKO tumor growth was comparable to the NTC controls (Supplemental Figure S3.2C). Similarly, end-point tumor weights were not significantly different between the two groups (Supplemental Figure S3.2D). These results demonstrate that TUTase loss does not impact tumor growth of KRASG12V transformed

NIH3T3 cells in the setting of an allograft model.

ZCCHC6 supports the growth and tumorigenicity in a subset of cancer cell lines

To determine if ZCCHC6/11 play a functional role in established human cancer cell lines, we took a loss-of-function approach. Initially, we focused on colon cancer because of the compelling evidence for TUTase overexpression in colon cancer cell lines and primary tumor samples (Figure

3.1A-D). First, we screened 14 gRNAs that target ZCCHC6 in the HCT116 colon cancer cell line using lentiviral constructs to achieve CRISPR/Cas9-mediated knockout. 6 out of 14 gRNAs yielded efficient ZCCHC6 knockout relative to two GFP-targeting controls (Supplemental Figure

S3.3A). Next, we compared the proliferation of HCT116 cells between two control GFP-targeting gRNAs, three inefficient ZCCHC6-targeting gRNAs and three highly efficient ZCCHC6-targeting gRNAs after 96 hours of growth. We found that only cells infected with the efficient ZCCHC6- targeting gRNAs exhibited reduced proliferation (Supplemental Figure S3.3B). Lastly, we performed triplicate experiments utilizing two efficient ZCCHC6-targeting gRNAs and observed a similar significant reduction in proliferation (Supplemental Figure S3.3C). We chose one

ZCCHC6-targeting gRNA (Z6KO) and knocked out ZCCHC6 in a larger panel of cancer cell lines.

Consistent with the results in HCT116 (Figure 3.2A), one additional cell line MCF7 (breast) that was also sensitive to ZCCHC6 loss when compared to controls (Figure 3.2B).

Given that ZCCHC6 and ZCCHC11 have been reported to redundantly uridylate RNA targets77,101,128, we surmised that some cancer cell lines may be only susceptible to double deletion. To test this hypothesis, we infected our cancer cell line panel with two gRNAs targeting each ZCCHC6 and ZCCHC11. One lung cancer cell line (H1299) exhibited decreased proliferation upon loss of both genes (Figure 3.2C), while many others were unaffected despite efficient knockout (data not shown). Furthermore, we found no correlation between sensitivity to

TUTase (either single or double) knockout and the presence of LIN28A/B expression, suggesting a LIN28-independent mechanism.

To determine if ZCCHC6 is required for tumorigenicity in vitro, we knocked out ZCCHC6 in

HCT116 cells and observed impaired anchorage-independent growth (Figure 3.2D). Then, we examined if ZCCHC6 loss reduces tumorigenicity in vivo using xenografts. To this end, we derived single cell HCT116 ZCCHC6 knockout clones, injected them subcutaneously into the flanks of immunocompromised (Rag2-/-gc-/-) mice and allowed them to grow for 45 days. We found that

ZCCHC6 loss led to decreased tumor growth, as indicated by reduced tumor volume and weight measurements (Figures 3.2H-K). Together, these results demonstrate that the TUTases support growth and tumorigenicity in a subset of cancer cell lines.

Figure 3.2. ZCCHC6 supports the growth and tumorigenicity in a subset of cancer cell lines. (A) Proliferation analysis of HCT116 cells after stable infection with CRISPR/Cas9 and either a non-targeting gRNAs (NTC) or highly efficient ZCCHC6-targeting gRNA (Z6KO), n=3. (B) Proliferation analysis of additional cancer cell lines sensitive to ZCCHC6 loss (n=3). Proliferation analysis of cancer cell lines sensitive (C) to ZCCHC6/11 (double knockout (DKO). (D) Soft agar assay. HCT116 cells were infected as in (A), plated in soft agar and grown for 3 weeks. Colonies were stained with crystal violet and counted, (n =3). (E-G) Western blot of ZCCHC6-targeting gRNAs used in (A)-(D). (H) Xenograft assay. HCT116 cells were injected into flanks of immunocompromised mice, (n =4). (I) Growth curve analysis, end-point tumor weight and size (J) of ZCCHC6 clonal knockout and control tumors. (K) Western blot analysis of ZCCHC6 of xenograft tumors. Note, error bars = s.e.m.; n.s. = non-significant P>0.05; *P < 0.05, **P < 0.01, ***P = 1.4 x 10-8 (regression analysis); (n=7).

To gain insight into the molecular mechanisms driving the phenotypes resulting from ZCCHC6 loss, we first addressed the contribution of miRNAs, given that ZCCHC6/11 have been shown to regulate both miRNA biogenesis and function (See Chapter 1)72,202. We utilized HCT116

DICEREx5∆ cells in which exon 5 of DICER has been knocked out resulting in a hypomorphic state where miRNA expression is extremely low222. We surmised that if the proliferative phenotype is

66 miRNA-dependent, depletion of ZCCHC6 in HCT116 DICEREx5∆ would not change proliferative rates. To test this, we used siRNAs to knockdown ZCCHC6 in HCT116 DICERWT and DICEREx5∆ cells. Surprisingly, ZCCHC6 depletion impaired proliferation to a similar degree in both cell lines indicating that the cellular proliferation phenotype is mechanistically independent of miRNA regulation (Supplemental Figure S3.4A).

ZCCHC6 regulates mRNA uridylation and turnover in cancer cells

In light of recent reports linking ZCCHC6/11 to mRNA decay through uridylation93,100,101,155, we wondered if ZCCHC6 also regulates mRNA uridylation and stability in cancer cells. To assess mRNA uridylation status upon ZCCHC6 depletion, we utilized TAIL-seq100, a transcriptome-wide method for identifying the 3’ ends of polyadenylated mRNAs (Figure 3.3A). While we were unable to achieve a sequencing depth that was sufficient to assess uridylation on a gene-by-gene basis, an issue that has been reported by others223, aggregation of all aligned reads revealed that mRNA uridylation of short (<25As) and very short polyadenylated (<15As) mRNAs were reduced upon

ZCCHC6 knockdown, indicating that ZCCHC6 is a major contributor to mRNA uridylation in

HCT116 cells (Figure 3.3B).

To determine if altered mRNA uridylation impacts steady-state mRNA levels, we performed standard poly(A)-selected RNA-seq, in HCT116 cells, 96 hours after ZCCHC6 knockdown. Given that mRNA uridylation has been associated with decreased mRNA stability100,101,155, we expected to see a skewing towards higher mRNA levels upon ZCCHC6 knockdown. However, we found that only ~11% of genes were differentially expressed between the knockdown and control conditions, with no bias towards up or down-regulation (Supplemental Figure S3.5A). Because our previous TAIL-seq results revealed preferential uridylation of mRNAs with short poly(A) tails, we postulated that our RNA-seq data set might be biased due to a poly(A) selection step used in library preparation. We therefore repeated the library preparation using ribosomal RNA depletion

Figure 3.3. ZCCHC6 regulates mRNA uridylation and turnover in cancer cells. (A) Schematic of TAIL-sequencing allowing for global analysis of 3’ mRNA ends. (B) Uridylation frequency stratified by poly(A) tail length as determined by TAIL-seq 96 hours after transfection of an siRNA targeting ZCCHC6 (SiZ6, bottom) or a non-targeting control (SiNC, top) in HCT116 cells. (C) Scatter plot of average half-lives (n=3) of individual genes determined by actinomycin-D treatment followed by RNA-seq 120 hours after transfection with a ZCCHC6 targeting siRNA or non-targeting control. (D) Box plot of average half-life of all genes after control or ZCCHC6 knockdown. (E) qPCR validation of actinomycin-D RNA-seq. (F) qPCR analysis of small-RNA half-lives upon ZCCHC6 knockdown. For all qPCR experiments, relative expression was calculated by normalizing to the expression of GAPDH. Note, error bars = s.e.m.; n.s = non-significant P>0.05; *P < 0.05, **P < 0.01, (n=3); TAIL-seq schematic is adapted from Chang et al. 2014. Molecular Cell. in place of poly(A) selection. Again, we observed a similar fraction of differentially expressed genes with no bias towards increased levels of expression. The high correlation between rRNA- depleted and poly(A)-selected RNA-seq suggests that ZCCHC6 depletion has a minimal effect on steady-state mRNA levels (Supplemental Figure S3.5B & C).

Given that our steady-state RNA-seq analysis provided little insight into changes in mRNA flux, we wondered if mRNA decay might instead be altered. To address this possibility, we

68 performed global mRNA half-life analysis by repeating knockdowns of ZCCHC6 in HCT116 cells and inhibiting transcription with actinomycin D for 0, 2 and 4hrs followed by rRNA-depleted RNA- seq. We were able to reliably assess the half-lives of around 1,400 mRNAs with high confidence.

Indeed, we found that ZCCHC6 depletion increased mRNA stability, with the vast majority of genes (1,076) exhibiting prolonged half-lives (Figure 3.3C), and the average half-life of all genes significantly increasing from 8 to 11 hours (Figure 3.3D). Next, we validated these findings by confirming altered mRNA half-lives of select targets using qPCR (Figure 3.3E). Finally, we also measured the half-lives of a set of small non-coding RNAs and found no difference upon ZCCHC6 depletion (Figure 3.3F) suggesting that the influence of ZCCHC6 on RNA stability may be restricted to mRNAs, at least in cancer cells. Together, these data show that ZCCHC6 controls mRNA uridylation and turnover in cancer cells.

ZCCHC6 impacts the cell cycle in cancer cells

To determine if certain pathways are preferentially altered by ZCCHC6 depletion, we performed gene ontology (GO) analysis of genes with significantly changed mRNA half-lives. This revealed an enrichment for pathways related to the cell cycle (Figure 3.4A). Upon closer analysis, we found that the mRNA half-lives of many histone genes and cell cycle regulators were increased

(Figure 3.4B & C). To determine if ZCCHC6 depletion altered the progression through the cell cycle, we knocked down ZCCHC6 in HCT116 cells and used propidium-iodide to track the cell cycle 5 days post-transfection. Similar to the impairment of S to M transition in yeast lacking the homolog of ZCCHC6, Cid180, we found modest prolongation of S phase upon ZCCHC6 knockdown. This finding is also consistent with a recent report of ZCCHC6 enhancing the degradation of histone mRNAs during S-phase of the cell cycle97.

Given these observations, we next wondered if ZCHCC6 expression was dynamically regulated throughout the cell cycle. To address this question, we synchronized HCT116 cells

Figure 3.4. ZCCHC6 impacts the cell cycle in cancer cells. (A) Gene ontology (GO) analysis of genes with significantly changed mRNA half-lives from Figure 3.3C. Changes in mRNA half-lives for (B) histones and (C) cell cycle regulators upon ZCCHC6 knockdown. (D) Propidium-iodide cell cycle analysis in HCT116 5 days post- transfection with a control (siNC) or ZCCHC6-targeting (siZ6) siRNA. (n=2). (E) Western blot analysis of ZCCHC6 and select cyclins from a 14hr time course after release from a double- thymidine block. Protein expression of ZCCHC6 relative to Actin is shown above in red. Estimated cell-cycle phases are indicated below. used a double-thymidine block and collected samples upon release. Western blot analysis of select cyclins allowed us to track the protein expression across different cell-cycle phases.

Significantly, ZCCHC6 protein expression increased two-fold expression across the S to G2/M transition (Figure 3.4E). Interestingly, this correlates closely with the rapid clearance of histone mRNAs that happens at the end of S-phase90,91. Collectively, these results imply that ZCCHC6 contributes to the regulation of the cell cycle by influencing the mRNA stability of critical cell cycle regulators and histones.

ZCCHC6 depletion sensitizes cancer cells to DNA damage

Histone abundance is tightly controlled at the transcriptional, post-transcriptional and protein level224. Given these quality control mechanisms, we wondered if dysregulated histone mRNA half-lives induced by ZCCHC6 loss were sufficient to alter histone protein levels. Indeed, the expression of the core histone proteins H2A, H2B, H3 and H4 were higher in the clonal ZCHCC6 knockout (Z6KO) HCT116 line relative to a GFP control (Figure 3.5A). While this increase in histone protein levels was relatively modest, this result is quite striking considering that others have had to synchronize cell populations to study the impact of TUTase-mediated uridylation on histone turnover95. Additionally, we found that complementation of the Z6KO HCT116 line with overexpression of either human (hZ6) or mouse ZCCHC6 (mZ6) reduced the expression of all four core histone proteins (Figure 3.5A). These data support the conclusion that ZCCHC6 regulates histone protein expression, likely through altered mRNA turnover.

Degradation of histones has been recently shown to be an important aspect of DNA-damage response pathways87 and altered histone expression can sensitize cells to DNA-damaging agents88,225. To explore the relationship between ZCCHC6 and the response to DNA-damage, we treated GFP and Z6KO cells with the widely used chemotherapeutic cisplatin, and then assessed two commonly assayed markers of DNA-damage response pathways: phosphorylation of

CHK2226 and the histone variant H2AX227. We found that cisplatin induced both phosphorylation of CHK2 and H2AX to a greater extent in Z6KO cells relative to controls suggesting that TUTase knockout might be more sensitive to DNA-damage (Figure 3.5B). Curiously, untreated Z6KO cells also exhibited increased γ-H2AX indicating that disrupted histone homeostasis, resulting from loss of ZCCHC6, might contribute to genome instability in the absence of exogenous DNA- damaging agents (Figure 3.5B).

To determine if TUTase loss sensitizes cells to DNA-damage, we assayed cell viability of GFP and Z6KO HCT116 cells treated with a range of cisplatin doses. We found that overall Z6KO cells

Figure 3.5. ZCCHC6 depletion sensitizes cancer cells to DNA damage. (A) Western blot analysis of core histone proteins in the following HCT116 clonal lines: control (GFP), ZCCHC6 knockout (Z6KO) or ZCCHC6 KO with re-expression of either human (hZ6) or mouse ZCCHC6 (mZ6) protein. (B) Western blot analysis of makers of DNA-damage response after treatment with cisplatin (65uM) for 48hrs in the indicated HCT116 cell lines. (C) Cell viability analysis of HCT116 GFP and Z6KO cell lines assessed with CellTiterGlow after 48 hrs of treatment with DNA-damaging agents (n= 3). (D) Pilot experiment (n=1) of cell viability after 48hrs of cisplatin treatment with the indicated HCT116 lines. Note, error bars = s.e.m.; n.s = non-significant P>0.05 *P < 0.05, **P<0.01, ***P<0.001. exhibited significantly reduced viability relative to GFP cells (Figure 3.5C). Similarly, we observed decreased viability of Z6KO when treated with three additional chemotherapeutics, whose mechanisms of action include DNA-damage: 5-flourouricil228, hydroxyurea229 and etoposide230

(Figure 3.5C). Finally, in a pilot experiment, we found that overexpression of either human (hZ6) or mouse (mZ6) ZCCHC6 partially rescued, while a catalytic-dead mutant of mouse Zcchc6

(mZ6DD) had no impact on the sensitivity of Z6KO cells to cisplatin treatment (Figure 3.5D).

Taken together, these data indicate that TUTase loss sensitizes cancer cells to a variety of DNA- damaging agents, possibly through disruption of uridylation-dependent control of histone protein homeostasis.

ZCCHC6/11 are dispensable for tumorigenesis in the APCmin model of colon cancer

Given our promising in vitro results for colon cancer cell lines and our expression data in primary colorectal tumors showing ZCCHC6/11 overexpression, we sought to further explore the role of the TUTases in vivo using the APCmin model of colon cancer. APCmin mice have a very short latency period before developing numerous intestinal and colonic tumors at around 4-6 weeks of age.231 Thus, this model allowed us to rapidly assess the in vivo role of the TUTases in tumorigenesis.

To determine if the TUTases are relevant to tumor formation in APCmin mice, we first interrogated ZCCHC6 and ZCCHC11 protein expression in colonic and intestinal tumors harvested from APCmin mice by Western blotting. Given previous work from our laboratory implicating LIN28B as an important oncogene in colon cancer, we also included tumors from two transgenic lines that overexpressed either LIN28A and LIN28B232 in the background of APCmin.

We found that ZCCHC6/11 protein expression is very low in normal intestinal and colonic mucosa, but high in tumors across all three APCmin lines (Supplemental Figure S3.6A). Additionally, LIN28A or LIN28B overexpression did not impact the level of TUTase expression suggesting LIN28- independent mechanisms. This was strikingly similar to the pattern in primary human colorectal tumor samples, in which 50% of tumors had high ZCCHC6/11, while normal mucosa samples

73 exhibited minimal levels of expression (Figure 3.1D). Thus, we conclude that the APCmin model recapitulates the overexpression of ZCCHC6/11 observed in primary human colorectal tumors.

To test the contribution of the TUTases to tumorgenesis in vivo, we obtained C57BL/6J mouse lines that harbor alleles of ZCCHC6 and ZCCHC11 in which critical exons were flanked by LoxP sites136. We then crossed these ZCCHC6/11 quad-flox mice with a C57BL/6J line with an intestinal specific Villin-Cre (B6.Cg-Tg(Vil1-cre)997Gum/J; Jackson Laboratories) to achieve a tissue restricted double TUTase knockout (DKO). We obtained the expected Mendelian ratios from this cross indicating that TUTase expression is not required for normal intestinal development (data not shown). Western blotting showing that ZCCHC6/11 were very efficiently depleted in all sections of the small intestine and colon from DKO mice validated our knockout strategy

(Supplemental Figure S3.6C). To generate intestinal specific DKO and control (FLOX) mice in the background APCmin, we followed the breeding scheme outlined in Supplemental Figure S3.6B.

We then analyzed the tumor burden in a cohort of 10 FLOX and 12 DKO mice at 17 weeks of age. Surprisingly, we observed no significant differences in the average tumor number per mouse,

(Supplemental Figure S3.6D) even when stratified by tumor size or anatomical location

(Supplemental Figure S3.6E). Additionally, we observed no obvious histologic differences between FLOX and DKO tumors (Supplemental Figure S3.6F). Interestingly, there was a trend towards DKO mice having a higher tumor burden particularly when considering large tumors.

However, this difference was not statistically significant. Therefore, we conclude that ZCCHC6/11 are dispensable for tumorigenesis in the APCmin model of colon cancer. This is consistent the observation that TUTase loss is dispensable for KRASG12V-mediated transformation of NIH3T3’s.

Together, these data indicate that TUTases are likely not required for oncogenic transformation

(Supplemental Figure S3.2). In contrast, our loss-of-function experiments in established cancer cell lines support a role for the TUTases in tumor maintenance (Figure 3.2).

3.3. Discussion.

Our in vitro and in vivo loss-of-function studies reveal that the oncogenic properties of some cancer cell lines are impaired by either the loss of ZCCHC6 or both ZCCHC6/11. Additionally, knockdown of ZCCHC6 alone was sufficient to decrease global 3’ mRNA uridylation and increase mRNA stability, including many cell cycle related genes and core histones. Furthermore, ZCCHC6 knockout sensitized cancer cells to DNA-damaging agents in a catalytic-dependent manner potentially through disrupted histone protein homeostasis. Collectively, these data implicate altered mRNA turnover as a previously unappreciated mechanism by which the TUTases are contributing to oncogenesis.

Our results expand the functional role of the TUTases in cancer well beyond the LIN28/let-7 axis. Profiling of human cancer cell lines uncovered prevalent TUTase expression in many contexts were LIN28A/B expression is absent indicating that the TUTases function independent of LIN28A/B in the majority of tumors. Our experiments using HCT116 DICEREx5∆ cells argue that impaired cellular proliferation induced by TUTase loss is miRNA-independent. This begs the question: what is the underlying mechanism driving changes in cellular proliferation? To address this, it would be interesting to test the importance of the catalytic activity of TUTases using similar biochemical approaches to those employed in Chapter 2. The sensitization of cells to DNA- damaging agents appears to be a uridylation-dependent phenomenon that is potentially driven by disrupted histone homeostasis. It is tempting to speculate that disrupted histone protein levels may also explain the altered proliferative capacity of ZCCHC6 depleted cancer cells given the known cytotoxic effects of excessive histones86. This could be directly tested by seeing if modest depletion of histones by siRNA knockdown rescues the proliferative impairment.

The proliferative and DNA-damage sensitization phenotypes induced by ZCCHC6 loss implicate the TUTases as potential therapeutic targets in cancer. However, it is not immediately clear which tumors would be susceptible. Some cancer cell lines did not exhibit a proliferative defect upon TUTase depletion, while others were susceptible to loss of ZCCHC6 alone or in

75 combination with ZCCHC11. This suggests that there are molecular differences between responding and non-responding tumor types. While our analysis of commonly perturbed pathways in cancer, (such p53 status and the expression of LIN28A/B or c-MYC) did not stratify cell lines by their responsiveness to TUTase loss (data not shown), this avenue of inquiry deserves additional effort.

Furthermore, it remains unclear to what extent there are functional differences between

ZCCHC6 and ZCCHC11. For example, it would be interesting to determine if ZCCHC11 depletion also sensitizes cells to DNA damage. While structurally quite similar, there are likely differences in the function of ZCCHC6 and ZCCHC11 across different cell lines given that ZCCHC11 was generally dispensable maintaining proliferative rates, while ZCCHC6 was not. This could potentially be explained by the relative expression level of each TUTase, the target RNAs and/or the presence of yet to be identified specifying cofactors acting similar to LIN28A/B.

Given the importance of the TUTases in a number of established cancer cell lines, it was surprising that the TUTases were dispensable for oncogenic properties of KRASG12V transformed

NIH3T3 cells. It would be interesting to try other oncogenic drivers such as c-MYC or mutant p53, or additional systems that are commonly used to model oncogenic transformation including the ba/f3 (pro-B cells)233 or MCF10A (human breast epithelia)234 cell lines. It would also be informative to explore the impact of the TUTases in the context of other oncogenic properties such as migration and invasiveness.

Our in vivo studies of TUTase function in the APCmin model of colon cancer yielded a puzzling result in that DKO mice trended towards a higher tumor burden relative to controls (Supplemental

Figure S3.5D & E). Given that this model requires the loss-of-heterozygosity of APC for tumor initiation235 and that Villin-Cre driven knockout almost certainly occurs before e13.0236, this model is an in vivo test of tumor initiation as opposed to tumor maintenance. This result, together with our observation that ZCCHC6 deletion can be deleterious to established colon cancer cell lines, is consistent with the growing appreciation that the genes involved in tumor maintenance and

76 oncogenic transformation are not always overlapping237,238. Additionally, there are examples of genes which can act both as oncogenes and tumor suppressers depending on the cellular context239,240. Therefore, it would be interesting to repeat these genetic experiments with an tamoxifen inducible Villin-cre236 to determine if the TUTases are required for tumor maintenance or progression in vivo by inducing knockout after tumor formation has occurred.

Taken together, our data suggest an oncofetal function for the TUTases particularly in the context of tumor maintenance, and implicate TUTase-dependent mRNA turnover as a previously unappreciated vulnerability of cancer. This highlights the therapeutic potential of TUTase inhibition particularly in contexts where DNA-damaging agents are utilized.

3.4. Methods.

Plasmids

Clonal NIH3T3 lines TUTase knockout lines were derived using CRISPR/Cas9 and gRNAs targeting mouse Zcchc6 or Zcchc11, or non-targeting control (NTC). gRNAs were cloned into the pSpCas9-2A-GFP or pSpCas9-2A-mCherry (See Chapter 2 methods for more detail). TUTase single and double knockouts cell lines were generated using population based infections of lentiviral CRISPR/Cas9 using gRNAs targeting human ZCCHC6 (Z6KO) or GFP (NTC) that were cloned into pLentiCRISPRv2-Puro. Double knockout lines (DKO) were generated by cloning

ZCCHC11 targeting gRNAs into pLentiGuide-Puro. gRNAs were designed as previously described (Chapter 2 Methods). Cell lines with stable overexpression of human ZCCHC6 were derived by cloning ZCCHC6 cDNA into pLentiCas9-Blast as previously described (Chapter 2

Methods). Overexpression of mutant KRAS was achieved using pBabe-Puro-KrasG12V. Kras- transformed NIH3T3 cells were generated by infected with retroviral particles produced as described in Chapter 2 methods with one exception; the following packaging plasmids used were used: retro-gag/pol and VSV.G. All plasmids were obtained as described: pLentiGuide-Puro

(Addgene plasmid #52963) and pLentiCRISPRv2-Puro (Addgene plasmid #52963), were gifts from F. Zhang (MIT). VSV.G (Addgene plasmid #14888) and retro-gag/pol (Addgene plasmid

#14887) were gifts from T. Reya (UCSD). pBabe-Puro-KrasG12V (Addgene plasmid#9052) was a gift from W. Hahn (DFCI). The gRNAs used in this chapter are as follows: NTC-1

GTAGGCGCGCCGCTCTCTAC, NTC-2 GGGCCCGCATAGGATATCGC, mZcchc6

GTGCTTATGAGCAAACGGAA, mZcchc11 AATCCGCCAGGACATTGTGG, GFP-1

GGGCGAGGAGCTGTTCACCG, GFP-2 GAGCTGGACGGCGACGTAAA, hZCCHC6

GTGGCTGTCATTCATCCAAG and hZCCHC11 TGTCCCAAGGATACCCGATT.

Cell Culture

HCT116 DICERWT (ATCC-CCL-247) and DICEREx5∆ (#HD R02-019), MCF7 (ATCC-HTB-22) and

U2OS (ATCC-HTB-96) lines were all cultured in DMEM supplemented with 10% FBS, 2mM glutamate, 1 U/ml Penicillin and 1 μg/ml Streptomycin. H1299 (ATCC-CRL-5803) cells were cultured similarly using an RPMI base. NIH3T3 cells (ATCC-CRL-1658) were grown in DMEM and 10% calf serum.

Protein lysates for Figure 3.1B were derived from the following cell lines and were loaded in on the gel in the same order from left to right:

Cancer type Cell line Culture medium Leukemia K562 DMEM:10%FBS MCF7 DMEM:10%FBS Breast MDA-MB-231 DMEM:10%FBS T47D RPMI:10%FBS Caco-2 DMEM:10%FBS HCT116 DMEM:10%FBS Colon LS174T DMEM:10%FBS RKO RPMI:10%FBS SW620 DMEM:10%FBS HepG2 DMEM:10%FBS Liver Huh6 DMEM:10%FBS Huh7 DMEM:10%FBS Lung A549 DMEM:10%FBS

H1155 DMEM:10%FBS H1299 DMEM:10%FBS H460 RPMI:10%FBS Fibroblasts BJ1 DMEM:10%FBS D283 EMEM:10%FBS Daoy EMEM:10%FBS Neural BE2C DMEM:10%FBS NBLS DMEM:10%FBS SK-N-FI DMEM:10%FBS M14 EMEM:10%FBS Skin SKMEL2 EMEM:10%FBS SKMEL28 EMEM:10%FBS A2780 DMEM:10%FBS Ovarian OVCAR3 DMEM:10%FBS SKOV3 DMEM:10%FBS CCG DMEM:10%FBS Kidney 293T DMEM:10%FBS Cervical HeLa DMEM:10%FBS

Primary Colorectal Cancer Tumor Samples

Colorectal cancer tumor samples were harvested from patients as previously described232. In all cases, informed consent was obtained in accordance with the guidelines approved by the human subject committees at the Brigham and Women’s Hospital and Harvard T.H. Chan School of

Public Health. APCmin tumor lysates were a gift from HC Tu, and were generated as previously described232.

APCMin Mouse Tumor Analysis

C57BL/6J mouse lines harbor floxed alleles of ZCCHC6 and ZCCHC11 were a gift from Dónal

O’Carroll and were genotyped as previously described136. ZCCHC6/11 quad-flox mice were crossed with a C57BL/6J line with an intestinal specific Villin-Cre (B6.Cg-Tg(Vil1-cre)997Gum/J;

Jaxson Laboratories) as indicated in the text. 17-18 weeks late mice were sacrificed and intestinal and colonic samples were harvested and analyzed using the “Swiss roll” method as described previously241. Paraffin embedded sections were stained with eosin and hematoxylin and then analyzed by light microscopy. Adenomas were identified by morphology and scored as previously

79 described232. All animal experiments were in accordance with guidelines approved by Boston

Children’s Hospital Institutional Animal Care and Use Committee.

Viral Production and Stable Cell Line Generation

Lentiviuses were produced as described in Chapter 2 and retroviruses were generated as previously described119. Stable cell lines were generated by transduction with unconcentrated viral supernatant and selection with 10 ug/ml Blasticidin or 1 ug/ml Puromycin, depending on the plasmid.

siRNA Transfections

Reverse transfections were carried out in 6 well plates using 1.5 x 105 HCT116 cells per well.

Each well was transfected with 40 pmol of siRNA with 5ul of Lipofectamine RNAiMAX as per the manufacturer’s instructions. (Invitrogen). The following siRNAs were utilized: siNC (Ambion

#4390843) and siZCCHC6 (Ambion s36058).

Knockout Cell Line Generation

To generate clonal-based CRISPR/Cas9 knockout, NIH3T3 double knockout clones were generated by transfecting pSpCas9-2A-GFP and pSpCas9-2A-mCherry plasmids with Zcchc6,

Zcchc11 and non-targeting control (NTC) gRNAs using Lipofectamine 2000 (Invitrogen). Double positive cells were FACS sorted and plated as single cells into a 96 well plate. TUTase expression was then assessed using Western blotting (See below). Population TUTase knockouts were generated by infecting cells with lentiviral particles derived from pLentiCRISPRv2-Puro then treating cultures with 1 ug/ml Puromycin for 1-3 days for selection of infected cells. To generate the H1299 DKO line, cells infected a second time with lentivirus generated from pLentiGuide-

Puro.

Cell Proliferation Analysis

Rates of cellular proliferation were determined by staining with trypan-blue counting live cells with a hemocytometer (Thermo Fisher Scientific # 22600100). All cancer cells were plated at a density of 5.0 x 105 cells/well on two 12-well plates, while NIH3T3’s at a density of 2.5 x 105 cells/well.

Day 0 counts were obtained 12-24hrs later to ensure equal starting numbers. The second plate was counted 96 hrs after Day 0 with cells being split as need to prevent reaching confluency.

Proliferative rates were calculated by multiplying 96hr counts by split ratios and then by dividing by Day 0 counts. Final proliferative values were calculated by normalizing to the appropriate controls.

Drug Treatments

HCT116 clonal control (GFP) or ZCCHC6 knockout (Z6KO) lines were plated in 96 well plates at a density of 2.5 x 104 cells/well. Cell were then treated for 48 hours with the following drugs at the concentrations indicated in the text: fluorouracil (5FU) (Fisher Scientific), hydroxyurea (Fisher

Scientific) and cisplatin (Sigma), etoposide (Fisher Scientific). Cell viability was then assessed using either CellTiter-Glo (Promega) or Cell Counting Kit-8 (Dojindo). To assess protein-level changes in DNA-damage markers, Western blotting was performed as previously indicated (see below; Chapter 2 methods) with the following exception: prior to lysis, phosphatase inhibitors

(Pierce) were added to RIPA buffer.

Soft Agar Assays

Soft agar assays were carried out as previously described242 with the following changes: Agarose

(Lonza) was prepared with growth medium to produce a 0.5% basal and 0.37% top layer. 0.5 x

103 HCT116 cells were plated in a well of a 12-well plate. Cultures were grown for 3 to 4 weeks

81 to allow colonies to arise. Growth media was changed every 2-3 day and colonies were then quantified using manually counting.

Allograft and Xenograft Assays

Allograft and xenograft assays were carried out as previously described242 with minimal changes.

Briefly, the flanks of Rag2-/-γc-/- mice (JAX #014593) were subcutaneously injected with 0.5 x 106

NIH3T3-KRASG12V or 1 x 106 HCT116 cells. Tumor growth was assessed using digital calipers.

Tumors were harvested 15-17 (NIH3T3-KrasG12V) or 41-43 (HCT116) days later. Boston

Children’s Hospital Institutional Animal Care and Use Committee approved all animal protocols and procedures utilized in this study.

Cell Cycle Analysis

HCT116 where synchronized using a double-thymidine block. Briefly, 2 mM thymidine (Sigma) was added to the culture media for 18. Cells were washed with PBS and allowed to recover in growth media for 8 hours. Cells were again treated with 2 mM thymidine 16 hours followed by one final media change. Samples were then collected as indicated. Cell cycle, was assessed by staining with propidium iodide (PI-BD Pharmingen) according to the manufacturer’s instructions.

Cells were then analyzed on the MACSQuant VYB (Miltenyi Biotec) and profiles were generated using the ModFit LT software.

Western Blotting

Normal adult human tissue lysates were purchased from Zyagen (San Diego, USA). Primary human colorectal tumors were obtained as described above and protein lysates were generated using the same protocol used to generate lysates from primary mouse tissues (Chapter 2

Methods). All protein lysates were analyzed as previously described. Secondary antibodies used

82 for chemiluminescence and fluorescence blots are listed in Chapter 2 methods. The following primary antibodies used in this chapter are listed below:

Antibody Catalog # Dilution anti-Actin Santa Cruz sc-1616 1:2,000 anti-α/β-Tubulin Cell Signaling #2148 1:1,000 anti-Gapdh Santa Cruz sc-32233 1:40,000 anti-ZCCHC6 Proteintech #25196-1-AP 1:1,000 anti-ZCCHC11 Proteintech #18980-1-AP 1:500 anti-Lin28a Cell Signaling #3978 1:1,000 anti-Lin28b Cell Signaling #4196 1:1,000 anti-cyclin A Santa Cruz sc-239 1:500 anti-cyclin B1 Santa Cruz sc-245 1:500 anti-cyclin D1 Santa Cruz sc-753 1:500 anti-H2A Abcam ab18255 1:1,000 anti-H2B Abcam ab1790 1:1,000 anti-H3 Active Motif # 39163 1:5,000 anti-H4 Abcam ab10158 1:1,000 γ-H2AX Abcam ab11174 1:1,000 anti-Chk2 Cell signaling #2662 1:1,000 anti-P-Chk2 Cell signaling #2197 1:1,000

Quantitative RT-PCR (qPCR)

Either total RNA or large (>200nt) RNA was isolated using Trizol (Invitrogen) and miRNeasy columns (Qiagen) according to the manufacture’s protocol. cDNA libraries were generated using

100-500 ng RNA and the miScript II RT kit (Qiagen). The hiFlex buffer system allowed for analysis of both mRNA and miRNAs from the same cDNA library. qPCR reactions were performed according to the manufacture’s recommendations using SYBR(Qiagen). Gene expression was calculated using the ΔΔCt method by normalization to either ACTIN or GAPDH. qPCR primer pairs were designed using the IDT PrimerQuest design tool (IDT) or were obtained from Zeisel et al.’s qPCR primer database208. The primers listed below were used in this chapter:

Primer Name Sequence hGAPDH-F ACCCAGAAGACTGTGGATGG hGAPDH-R TTCAGCTCAGGGATGACCTT hZCCHC6-F TCCAGTTTCCAGCCATTATGT hZCCHC6-R CTAGCATGGAAGTCTGCATCA

hZCCHC11-F CGACACATCTGCTACCTCTTG hZCCHC11-R TACTATCGTTGGTGGCTTGC hHMGA2-F GACCTAGGAAATGGCCACAA hHMGA2-R ACTGCTGCTGAGGTAGAAATC hMTF2-F CTAGATGGTCAGATGGCTTGTT hMTF2-R AGTGGCTCCTGTTTGAATGT hENTPD7-F GCATTCCTGGGACTCTTCTT hENTPD7-R CGAGCCAAATACCTTTCGTATTG hTLDC1-F AGCTGCAAGATGGCAAGA hTLDC1-R ACGAGCACAGGATGAGAAAG U6 Qiagen (MS00033740) miR-154 Qiagen (MS00003598) miR-211 Qiagen (MS00003808) miR-382 Qiagen (MS00031836) Let-7a Qiagen #MI0000556 Let-7b Qiagen # MI0000558 Let-7c Qiagen # MI0000559 Let-7e Qiagen # MI0000561 Let-7f Qiagen # MI0000562 Let-7g Qiagen # MI0000137

RNA Sequencing

Total RNA (100-500ng) of RNA was enriched with either poly(A) selection (NEBNext Poly(A) mRNA Magnetic Isolation Module) or rRNA depletion (Ribo-Zero-Gold; Illumina). Libraries were constructed using NEBNext Ultra RNA Library Prep Kit. Average library size was estimated using either a Bioanalyzer (Agilent) or 4200 Tapestation (Agilent). Library concentration was determined using both qRT-PCR (Kapa Biosystems) and Qubit dsDNA high-sensitivity assay (Invitrogen).

Sequencing was performed on either the NextSeq 500 or HiSeq 2500 platforms (Illumina) using

75-bp single-end reads. Gene expression (RPKM) was determined with TopHat209 and HTSeq- count210. The DAVID 6.8 Functional Annotation Tool was use for GO analysis189,190.

TAIL-Sequencing

TAIL-seq libraries were generated and analyzed as previously described100,101. However, poly(A) spike-in controls were omitted. Instead, poly(A)-tail length were estimate standard base-calling

84 algorithms and binned into categories (<15nt, <25nt, and ≥25) (Illumina). A detailed protocol is available upon request.

RNA half-life analysis

HCT116 cells were transfected with either a control or ZCCHC6-targeting siRNA as described above. Cells were then cultured for 96 hours prior to adding actinomycin-D (Themo scientific-

11805017). Actinomycin-D treatment, RNA-sequencing and analysis allowing for the determination of RNA half-lives were conducted as previously described101.

Statistics

Statistical significance of P<0.05, P<0.01 or P<0.001 are denoted with one, two or three asterisks respectively. P values were calculated using a two-tailed (unless otherwise specified) using

Student’s t-test. Data are shown as mean of biological replicates with error bars displaying variation as calculated by s.e.m. In the case of RNA half-life measurements, paired t-tests were used and the Benjamini-Hochberg correction was applied to account for multiple hypotheses testing.

Data Availability

All data contained within this manuscript is available upon request. RNA-sequencing data sets will be deposited in the Gene Expression Omnibus (GEO) prior to publication.

AUTHOR CONTRIBUTIONS

This chapter represents a manuscript in preparation co-authored by D.S. Pearson, K.M. Tsanov and Y.C. Huang. D.S. Pearson and K.M. Tsanov designed and performed most of the experiments for Figures 3.1-4, while Y.C. Huang performed the experiments for Figure 3.5 under

85 the guidance of D.S Pearson and K.M. Tsanov. Specific contributions are as follows: D.S Pearson performed most of the cloning and generated the text Figures 3.1C & D; 3.2A, C, E & F; Figure

3.3; Figure 3.4 A-C; Supplemental Figure S3.1; Supplemental Figure S3.2A & B; Supplemental

Figure S3.5 & Supplemental Figure S3.6. K.M. Tsanov generated Figures 3.1A & B; 3.2B, D-J;

3.4D& E; Supplemental Figure S3.2; Supplemental Figure S3.3C & D; and Supplemental Figure

S3.4. and portions of the Methods section. Y.C. Huang and J. Barragan helped with cloning,

APCmin tumor counting, and Western blot analysis. A. Han, L. Pantano, J. Han and performed

RNA-seq analysis. H.C. Tu, Z. Qian, S. Ogino and C.S. Fuchs donated primary human tumor samples. J.K. Osborne and J.T. Powers helped with experimental design. O. Hofmann supervised

TAIL-seq bioinformatics analysis. G.Q. Daley supervised the study and edited the manuscript.

ACKNOWLEDGMENTS

We would like to thank S. Buratowski, J. Rinn, R. Horvitz, and the members of the Daley lab for many insightful discussions. Additionally, we thank Dana-Farber/Harvard Cancer Center in for the use of the Rodent Histopathology Core, which performed paraffin embedding, sectioning and H

& E staining. Dana-Farber/Harvard Cancer Center is supported in part by a NCI Cancer Center

Support Grant # NIH 5 P30 CA06516.R. Thanks to R. Mathieu and M. Paktinat for helping with flow cytometry. Thank you to R. Rubio CCCB (DFCI) and X. Wu/Yi Zhang laboratory for technical help in performing the RNA-seq. IDDRC Molecular Genetics Core (BCH) performed the

Bioanalyzer analysis and was supported by NIH (NIH-P30-HD 18655). Intensive informatics analysis was performed utilizing the Orchestra High Performance Computing Cluster (HMS). D.S.

Pearson is supported by a grant from NIGMS (T32GM007753). K.M. Tsanov was a Herchel Smith graduate fellow and a Howard Hughes Medical Institute international student research fellow.

G.Q. Daley was supported by the Howard Hughes Medical Institute and a grant from the NIGMS

(R01GM107536). G.Q. Daley also is an investigator of the Manton Center for Orphan Disease

Research.

CHAPTER 4

Discussion and Perspective

4.1. TUTases are regulators of cell state transitions.

In recent years, 3’ RNA uridylation has emerged as an important epitranscriptomic mark that contributes to the regulation of gene expression through multiple mechanisms. However, the biologic functions of many of the “writers” of terminal uridylation, including ZCCHC6 and

ZCCHC11, remain poorly understood. The work presented in this dissertation addresses this knowledge gap by uncovering novel functions of these TUTases in normal development and oncogenesis. This suggests an overarching role for TUTases and RNA uridylation in guiding cell state transitions.

Cell state transitions are defined by changes in gene expression that give rise to distinct phenotypic properties243. These transitions are at the heart of developmental programs as well as adaptation to environmental stress including hypoxia, nutrient deprivation and drug treatment244,245. In addition to our observations that the TUTases influence the differentiation of mESCs and muscle progenitors, the few previously reported phenotypes support a role for the

TUTases as regulators of cell state transitions. For example, in mouse and human lung cell lines, the importance of Zcchc11 in regulating cytokine elaboration is uncovered upon exposure to inflammatory stimuli72. Similarly, whole body knockout of Zcchc6 resulted in no overt phenotypes in mice156; yet, upon exposure to S. pneumoniae, defects in innate immunity became apparent.

Additionally, both TUTases are required for one of the earliest cell state transitions, oogenesis136.

This may be a common feature of uridylation-based mechanisms, whereby their function is particularly relevant to transitions rather than steady states. Consistent with this, unpublished results demonstrate that Dis3l2 knockout mESC exhibit no overt phenotypes in serum/LIF culture conditions. However, upon differentiation into embryoid bodies, Dis3l2 knockout cells display an increased propensity toward renal lineages (Pirouz M. et al. personal communication).

It would be interesting to examine additional cell state transitions to determine if there are particular types that are preferentially sensitive to TUTase activity. Embryonic development is an

88 obvious starting point because it is the result of numerous carefully coordinated cell state transitions that require the integration of many cell autonomous and non-autonomous signals.

The fidelity of developmental timing is known to result from the integration of many molecular mechanisms, with post-transcriptional regulators playing a prominent role246. Therefore, TUTases most likely contribute to embryogenesis, at least in part, by ensuring that the timing of critical cell fate switches occurs with high precision.

Additionally, somatic cells can be reprogrammed to a pluripotent state, albeit at low efficiency, through overexpression of defined factors247. This reprogramming process is a complex multi- stage journey that involves numerous cell state transitions248. This would be an interesting context to assess the contribution of the TUTases, as detailed molecular characterization has revealed that reprogramming recapitulates many aspects of development “in reverse”249. Given that high

TUTase expression correlates with less differentiated cell states, and that enforced expression impairs differentiation, it is tempting to speculate that TUTase overexpression may enhance the efficiency of somatic cell reprogramming. On the other hand, if TUTase activity is an overall barrier to cell state transitions, overexpression would instead impair reprogramming. It would be interesting to differentiate between these models, which could yield further insights into the cell fate regulatory function of the TUTases.

Akin to reprogramming, tumorigenesis has long been conceptualized as multi-step process that progresses through defined and qualitatively different intermediate cell types250. This progression model was pioneered in colorectal cancer, where mutations in genes controlling cellular differentiation and growth accumulate over time giving rise to a four-step progression to neoplasia251. Conceptualizing tumorigenesis as a series of cell state transitions provides an explanation for our curious observation that tumor burden was higher in the TUTase DKO APCmin mouse model, which we examined in Chapter 3. This would be consistent with the hypothesis that TUTases indeed provide a barrier to cell state transitions relevant to the neoplastic progression. Given that the difference between the tumor burden of control and DKO animals

89 appeared modest, further studies with an expanded cohort size would be warranted.

Nevertheless, these findings provide a strong foundation for the interrogation of TUTase function in genetically defined in vivo models of cancer.

4.2. Uridylation-dependent vs. independent functions of the TUTases.

Previously reported functions of ZCCHC6 and ZCCHC11 have been mostly attributed to uridylytransferase activity. However, catalytic-independent roles have also been uncovered, highlighting the importance of addressing both possible mechanisms of action153,154. In Chapter

3, we demonstrate that ZCCHC6/11 support the proliferation and tumorigenicity of a subset of cancer cell lines and that ZCCHC6 loss increases their sensitivity to DNA damaging agents.

These phenotypes are most likely catalytic-dependent due to observed dysregulated mRNA half- lives of histones and cell cycle regulators (Figure 3.4). More definitively, the enhanced susceptibility to DNA damaging agents upon ZCCHC6 knockout can be rescued by wild type, but not catalytic-dead ZCCHC6 (Figure 3.5). Interestingly, our data implicating ZCCHC6 as a regulator of histone homeostasis, together with reports that disrupted histone mRNA decay can promote genome instability252,253, suggests a potential mechanistic model to explain the increased tumor burden in APCmin mice. The APCmin model recapitulates aspects of multi-stage tumorigenesis with a critical step being loss of heterozygosity (LOH) of APC254. Due to increased genome instability resulting from histone dysregulation, tumorigenesis could be enhanced through increased frequency of various genetic alterations including APC LOH. This hypothesis could be tested by comparing the genomes of tumors isolated from wild type and TUTase DKO

APCmin mice with the expectation that DKO tumors would harbor higher levels of genomic lesions, including copy number alterations.

In Chapter 2, we provide compelling evidence, using both point and truncation mutagenesis, that Zcchc6 regulates myogenesis through a uridylation-independent mechanism(s). While we were able to define an N-terminal domain of Zcchc6 that is required for its control of myogenesis,

90 it remains unclear how this protein domain regulates differentiation. There are many examples of catalytic-independent functions across diverse enzyme classes including, kinases255, phosphatases256, DNA-methyltransferases257, and deubiquitinases258. Non-enzymatic functions commonly include allosteric regulation of other enzymes, regulation of transcription, and structural functions like protein scaffolding. As the N-terminus of Zcchc6 contains a C2H2 zinc-finger domain that could play a role in facilitating TUTase-RNA interactions, we favor a model whereby the

TUTases can function as classic RNA-binding proteins. To address this possibility, we attempted to determine the direct RNA targets of the TUTases. We utilized two different RNA- immunoprecipitation (RIP) strategies: native-RIP and individual-nucleotide resolution in vivo UV- crosslinking and immunoprecipitation (iCLIP)259. Despite achieving efficient Zcchc6 pulldown at the protein level, we recovered little RNA with low enrichment above controls (data not shown).

This suggests that either Zcchc6-RNA interactions may be transient and/or weak, necessitating additional optimization of our immunoprecipitation protocols. The latter is further warranted given that both ZCCHC6 and ZCCHC11 were identified using an unbiased proteomics approach that employed human mRNAs as “baits”, indicating that TUTase-mRNA interactions occur260.

In addition to facilitating protein-RNA interactions, the zinc finger domains can also mediate

DNA binding or protein-protein interactions151,152, raising the possibility that TUTases may act as transcriptional regulators and/or protein scaffolds. In this regard, comprehensive identification of

TUTase protein-protein interactions through mass spectrometry would provide important insights into its non-enzymatic functions. Ideal for that purpose would be a comparison of Zcchc6 truncation mutants with and without an impact on muscle differentiation, which would directly point to functional relevance and readily enable its validation.

Distinguishing between the relative importance of catalytic-dependent and independent activities will be a critical component of any future investigations of TUTase function. With respect

91 to in vivo studies of TUTase function, the recent availability of mice harboring catalytic-dead alleles for both Zcchc6 and Zcchc11 will certainly accelerate such efforts136.

4.3 Regulation of the cell cycle: a unifying theme?

As discussed above, there is compelling evidence that the TUTases’ impact on muscle differentiation is uridylation-independent, while the cancer phenotypes we uncovered are likely dependent on uridylation activity. Is it possible that these seemingly disparate functions are mechanistically linked?

Cid1, the yeast homolog of ZCCHC6, regulates the S-M cell cycle checkpoint in S. pombe80.

Similarly, our cancer data indicate that ZCCHC6 depletion results in prolonged S-phase accompanied by impaired cell proliferation. Interestingly, exit from the cell cycle is an important, but not sufficient, step of many differentiation programs across a range of cell types261. This is particularly true for terminal differentiation systems including the in vitro models of skeletal muscle

187,262 differentiation we employed . In proliferating myoblasts, p21, a critical regulator of the G1 checkpoint of the cell cycle, is not expressed. However, upon serum withdrawal, there is irreversible upregulation of p21, which then inhibits the activity of cyclin-dependent kinases and causes cell cycle exit187. While the functional relationship between cell cycle exit and the transcriptional changes associated with differentiation is not fully understood, establishment of a post-mitotic state is required for the differentiated phenotype of skeletal muscle cells. Curiously, p21-/- mice develop normally263, but injury-induced muscle regeneration is delayed264,265.

Collectively, the above observations suggest that the TUTases’ differentiation roles we have discovered may also be mediated through regulation of cell cycle exit. While we did not observe a proliferative difference upon either knockdown or OE of the TUTases in undifferentiated myoblasts, it is possible that they affect the kinetics of cell cycle exit upon exposure to differentiation signals. Similarly, it would be interesting to interrogate the regenerative capacity of

92 skeletal muscles in Zcchc6 and Zcchc11 knockout mice upon injury, given that neither model exhibits overt skeletal muscle phenotypes in unperturbed conditions132,156.

4.4. Therapeutic implications of TUTase modulation.

The functional roles we uncovered for ZCCHC6/11 in cellular differentiation (Chapter 2) and cancer (Chapter 3) highlight many potential applications for TUTase inhibition including tissue regeneration and novel chemotherapeutic strategies for cancer. Specifically, our results demonstrating that TUTase depletion facilitates myogenic differentiation suggest that TUTase inhibition might be therapeutically exploited to enhance skeletal muscle regeneration in response to injury. Furthermore, excessive loss of differentiated skeletal muscle is a poor prognostic indicator for muscular dystrophies and myopathies as well as many systemic diseases including cancer-induced cachexia, diabetes and chronic obstructive pulmonary disorder266. Therefore,

TUTase inhibition maybe useful avenue to generally enhance muscle differentiation and regeneration in those contexts. It is likewise possible that TUTase inhibition would be useful for promoting the differentiation other clinically relevant tissues, which is an area that warrants further study.

In cancer, the development of TUTase inhibitors as a means of perturbing LIN28 function is an area of active investigation267. Our data demonstrating LIN28-independent roles of the

TUTases in cancer indicate these compounds might be beneficial in LIN28-negative tumors, extending their potential utility to a broader range of cancers. Furthermore, our observations that

TUTase loss sensitizes cells to DNA-damaging agents that are commonly used as chemotherapeutics suggests that TUTase inhibition may be particularly effective when used in combination with such agents and provide the basis of novel combination therapy regimens.

Further, impaired differentiation is thought to play a critical role in the etiology of numerous cancers including rhabdomyosarcoma268, acute promyelocytic leukaemia269 and

93 neuroblastoma270. An effective therapeutic strategy for many of these malignancies is to induce differentiation with small molecules like retinoic acid and arsenic271,272. Given our observation that

TUTase depletion enhances differentiation across multiple cell types, it would thus be a great interest to interrogate the potential synergy of these differentiation therapies with TUTase inhibition.

Of note, ZCCHC6/11 have also been implicated as facilitators of apoptosis through promotion of global mRNA decay in a uridylation-dependent manner155, raising the possibility that in some contexts the TUTases might be acting as tumor suppressor genes. Our in vivo data in the APCmin model are consistent with such a function, highlighting that context should be carefully taken into consideration in the potential application of TUTase inhibition-based therapies.

Finally, current screening strategies to inhibit the TUTases with small molecules have been focused on targeting its catalytic site267. However, our data in developmental systems indicate that screening for compounds that perturb the function of the N-terminus of ZCCHC6 rather than its catalytic domain would be similarly important. Overall, our results uncover novel functional roles for ZCCHC6/11, with therapeutic implications for both tissue regeneration and cancer.

AUTHOR CONTRIBUTIONS

This chapter is unpublished. D.S. Pearson wrote the text with edits from P. Missios, K.M. Tsanov,

T.E. North, and G.Q. Daley.

APPENDIX Supplementary Material

Supplemental Figure S2.1. ZCCHC6/11 regulate myoblast differentiation. (A) Western blot analysis of Lin28a expression in mESCs (V6.5) and differentiating primary myoblasts. (B) Proliferation analysis of C2C12 cells after transfection with control or Zcchc6/11 targeting siRNAs after 96 hours. Cells were transfected one day prior to plating for proliferation assays with either control (NC), Zcchc6 (Z6), Zcchc11 (Z11) or both (Z6/11) targeting siRNAs. (C) Proliferation analysis of empty vector (EV) and Zcchc6 wild type (Z6) or catalytic null (Z6DD) overexpressing C2C12 cells 96 hours after plating. (D) Western blot analysis of TUTase expression 48hrs after transfection of C2C12s with 6 different siRNAs targeting either Zcchc6 (Z6-1-3) or Zcchc11 (Z11-1-3) or a control siRNA (NC). (E) Bright-field images of C2C12 cells transfected with the corresponding siRNAs in (C) at day three of differentiation. (F) qPCR analysis of mature let-7 expression across C2C12 differentiation. Relative expression was calculated by normalizing to the expression of the small RNA U6. Note, n.s. = non-significant p>0.05; error bars = s.e.m.

Supplemental Figure S2.2. TUTase overexpression blunts the induction of differentiation genes.

(A) Log2 expression (RPKM) of genes at day minus one in EV relative to Z6 or (B) Z6DD overexpressing C2C12 lines. (C) Unsupervised hierarchal clustering of significantly changed genes between EV day minus one and day two across EV, Z6 and Z6DD lines. (D) Scatter plot comparing Log2 fold change relative to Z6 or Z6DD of significantly changed genes between EV and Z6DD at day two of differentiation. (E) Relative expression between EV, Z6 and Z6DD of the top 25 “differentiation” genes at day two of differentiation. (F) Gene ontology analysis of genes significantly upregulated in Z6 or (G) Z6DD relative to EV.

Supplemental Figure S2.2 (Continued)

Supplemental Figure S2.3. TUTase overexpression modestly impacts mRNA half-lives in proliferating C2C12s. (A) Schematic of experimental design for assessing mRNA half-life in proliferating C2C12s. (B) Western blot analysis of Zcchc6/11 expression across C2C12 differentiation. Scatter plot of average mRNA half-lives of individual genes determined by actinomycin-D treatment followed by RNA-sequencing comparing EV and either (B) Z6 or (C) Z6DD lines. Venn diagrams showing the overlap of genes between Z6 and Z6DD with significantly (D) decreased and (E) increased mRNA half-lives relative to EV. (n=3).

Supplemental Figure S2.4. TUTase overexpression does not influence the expression of pluripotency markers under steady-state conditions. (A) qPCR analysis of pluripotency markers in iZ6/11 v6.5 mESCs cultured with or without doxycycline (dox) for 72 hours in serum/LIF or (B) 2i/LIF. (C) Proliferation analysis of dox- inducible ZCCHC6 (iZ6) and ZCCHC11 (iZ11) overexpression mESC lines cultured in 2i/LIF. Cells were pre-treated with dox for five days and then proliferation was assessed 72 hours later. Note, error bars = s.e.m.; n.s = non-significant P>0.05; *P < 0.05, **P < 0.01 (n=3)

Supplemental Figure S3.1. Genetic lesions of ZCCHC6 and ZCCHC11 in human tumors. (A) Frequency of deletions (blue), amplifications (red) or no alterations (gray) of the ZCCHC6 and ZCCHC11 loci across 170 studies representing a total of 42,587 primary patient samples. The prevalence of patients with either an amplification or deletion are shown as a percentage of all patients next to the gene name. Note that each bar represents a single patient. Study of origin legend is available at http://www.cbioportal.org. (B) Distribution of point mutations and small insertions/deletions identified in ZCCHC6 and (C) ZCCHC11 is overlaid on a schematic of the protein domain structures. The y-axis indicates the number of samples harboring a given mutation, while the type of mutation is represented by a colored circle. All data was obtained from publicly available data sets from C-BioPortal. PAP = poly(A) polymerase.

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Supplemental Figure S3.2. ZCCHC6/11 are dispensable for NIH3T3 transformation by KRASG12V . (A) Western blot analysis of NIH3T3 clones derived using transient expression of Cas9 and either two control non-targeting gRNAs (NTC) or two gRNAs targeting ZCCHC6/11 (DKO). (B) Proliferation analysis of a representative NTC and DKO clone (n=5). (C) Transformed NIH3T3 cells were injected into flanks of immunocompromised mice (n=9). (D) Growth curve analysis and (E) end-point tumor weight of NTC and DKO allograft tumors. Note, error bars = s.e.m.; n.s = non-significant P>0.05.

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Supplemental Figure S3.3. CRISPR/Cas9-mediated deletion of ZCCHC6 in HCT116 cells. (A) Western blot screen of ZCCHC6-targeting gRNAs used in (B) and (C). (B) Proliferation analysis of HCT116 cells after CRISPR/Cas9-mediated knockout of ZCCHC6 with two GFP- targeting gRNAs (dark blue), three inefficient ZCCHC6-targeting gRNAs (light blue) or three highly efficient ZCCHC6-targeting gRNAs (dark red) (n=1). Cells were infected with Cas9/sgRNA lentivirus, selected with puromycin for 48 hours, then plated at equal cell numbers at day zero and re-counted 96 hours later. (C) Proliferation analysis of HCT116 cell infected with GFP-targeting (GFP1 or GFP2) or ZCCHC6-targeting (Z6_4 or Z6_8) lentivirus (n= 3). Day zero counts are shown in light gray. For statistical analysis, t-tests compared GFP1 to either Z6_4 or Z6_8. Note, error bars = s.e.m.; *P < 0.05,

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Supplemental Figure S3.4. ZCCHC6 impacts cancer cell growth independent of miRNAs. (A) Proliferation analysis 120hrs post transfection with either a control (siNC) or ZCCHC6 (siZ6) targeting siRNA in DICERWT and DICEREx5∆ HCT116 cells. Note, error bars = s.e.m.;**P<0.01.

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Supplemental Figure S3.5. Depletion of ZCCHC6 minimal alters steady-state mRNA expression. (A) Distribution of changes in mRNA expression 120 hours after ZCCHC6 knockdown in HCT116 cells using either poly(A) selection or (B) ribosomal RNA depletion (RiboZero) RNA- seq library protocols. (C) Correlation of Log2 fold changes after ZCCHC6 knockdown of individual genes between the two different methods of library preparation.

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Supplemental Figure S3.6. ZCCHC6/11 are dispensable for tumorigenesis in the APCmin model of colon cancer. (A) Western blot analysis of normal and tumor lysates from the indicated murine models of colorectal cancer. Each lane represents an independent tissue or tumor sample. (B) Breeding scheme used to generate intestinal/colon specific conditional deletion of Zcchc6 and Zcchc11 in the background of APCMin model of colon cancer. (C) Western blot analysis of Zcchc6 and Zcchc11 from Flox (Zcchc6fl/fl; Zcchc11fl/fl) and DKO (Villin-Cre; Zcchc6-/-; Zcchc11-/-) Mice. Duo = duodenum , jej = jejunum, ile = ileum, and col = colon. (D) Average tumor burden per Flox (n = 10) and DKO (n =12) mice. (E) Average tumor burden stratified by tumor size and anatomic location. Tumor size; S = small; less than one villus and L= large; greater than one villus. (F) Representative bright field images of hematoxylin and eosin stained intestinal tumors from Flox and DKO mice at low (Top) and high (Bottom) magnification. Red arrow heads mark tumors at low magnification, while dashed red lines circumscribe tumors at high magnification. Note, error bars = s.e.m.; n.s = non-significant.

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Supplemental Figure S3.6 (Continued)

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