Differential functions of mammalian Cdk7 and CCRK kinases in regulating transcription and ciliogenesis Ying Yang Institute of Biotechnology and Division of Genetics Department of Biosciences Faculty of Biological and Environmental Sciences and Helsinki Graduate Program in Biotechnology and Molecular Biology University of Helsinki ACADEMIC DISSERTATION To be presented with the permission of the Faculty of Biological and Environmental Sciences of the University of Helsinki for public examination in the lecture hall 2 of Infocenter Korona, Viikinkaari 11, Helsinki on June 13th, 2014, at 12 o'clock noon. HELSINKI 2014 Supervisor Tomi P. Mäkelä, MD., PhD., Professor Institute of Biotechnology University of Helsinki Helsinki, Finland Thesis follow-up committee Lea Sistonen, PhD., Professor Department of Biosciences Abo Akademi University Turku, Finland Reviewers Lea Sistonen, PhD., Professor Department of Biosciences Abo Akademi University Turku, Finland Päivi Ojala, PhD., K. Albin Johansson Research Professor Institute of Biotechnology University of Helsinki Helsinki, Finland Markku Varjosalo, PhD., Docent Institute of Biotechnology University of Helsinki Helsinki, Finland Opponent Jeremy Reiter, MD., PhD., Associate Professor Department of Biochemistry University of California, San Francisco California, USA Custos Minna Nyström, PhD., Professor Department of Biosciences University of Helsinki Helsinki, Finland ISBN: 978-952-10-9978-6 (paperback) ISBN: 978-952-10-9979-3 (PDF; http://ethesis.helsinki.fi) ISSN: 1799-7372 Painosalama Oy, Turku, Finland 2014 “If you thought that science was certain - well, that is just an error on your part.” -Richard P. Feynman TABLE OF CONTENTS LIST OF ORIGINAL PUBLICATIONS ................................................................................................... ii ABBREVATIONS ..................................................................................................................................... iii ABSTRACT ................................................................................................................................................ 1 REVIEW OF THE LITERATURE ........................................................................................................... 2 1. Eukaryotic RNA polymerases ........................................................................................................................... 2 2. RNA Pol II-‐mediated synthesis of mRNA ...................................................................................................... 3 2.1. The unique carboxy terminal repeat domain (CTD) of RNA Pol II .................................... 3 2.2. Role of chromatin structure in transcription ............................................................................. 4 2.3. The RNA polymerase II transcription cycle and CTD phosphorylation .......................... 4 2.4. Mutual regulation of transcription and decay ............................................................................ 7 2.5. Transcriptional cyclin-‐dependent kinases (Cdks) phosphorylate Pol II CTD ........... 10 2.6. Cdk7-‐cyclin H-‐Mat1 as the kinase subcomplex of TFIIH in general transcription .. 13 2.7. Cdk7-‐cyclin H-‐Mat1 in cell type-‐specific transcription ....................................................... 14 3. RNA polymerase I-‐mediated synthesis of rRNA ..................................................................................... 15 3.1. The structure of rRNA genes ........................................................................................................... 15 3.2. Pol I transcription processes on rDNA ....................................................................................... 16 3.3. Involvement of modifiers of Pol II CTD in rDNA transcription ........................................ 18 4. Regulation of ciliogenesis and links to cell cycle control .................................................................. 19 4.1. The structure of primary cilia ........................................................................................................ 19 4.2. IFT: the core machinery for cilium assembly and maintenance ..................................... 22 4.3. Bidirectional crosstalk between ciliogenesis and the cell cycle ...................................... 28 4.4. Cilium formation and length control ........................................................................................... 29 4.5. The cilia-‐mediated signaling and links to cancer ................................................................... 37 AIMS OF THE STUDY ........................................................................................................................... 41 MATERIALS AND METHODS ............................................................................................................. 42 RESULTS .................................................................................................................................................. 52 1. Cdk7-‐cyclin H-‐Mat1 kinase subcomplex of TFIIH is required for global transcription and mRNA turnover (I, II) ............................................................................................................................................. 52 2. Cdk7 and the associated Mat1 inhibits adipogenic transcription program through inhibitory phosphorylation of PPARγ (III) ................................................................................................... 54 3. Inhibition of ciliogenesis by CCRK and its effectors ICK and MAK promotes cell proliferation (IV) ..................................................................................................................................................... 55 DISCUSSION & PERSPECTIVES ......................................................................................................... 58 1. Cdk7 subcomplex of TFIIH in transcription mediated by RNA Pol II and Pol I ........................ 58 2. Crosstalk between RNA synthesis and degradation: an emerging layer of regulation for gene expression ........................................................................................................................................................ 59 3. Cell type-‐specific functions of Cdk7 subcomplex ................................................................................... 60 4. The role of CCRK and its substrates ICK/MAK in regulating ciliogenesis .................................. 61 5. Cilia, cell cycle and cancer .............................................................................................................................. 62 CONCLUSIONS ....................................................................................................................................... 64 ACKNOWLEDGEMENTS ...................................................................................................................... 65 REFERENCES .......................................................................................................................................... 67 i LIST OF ORIGINAL PUBLICATIONS This thesis is based on the following original publications, which are referred to in the text by Roman numerals: I. Helenius K, Yang Y, Tselykh T.V, Pessa H.K.J., Frilander M., Mäkelä T.P. Requirement of TFIIH kinase subunit Mat1 for RNA Pol II C-terminal domain Ser5 phosphorylation, transcription and mRNA turnover. Nucleic Acids Res. 2011 Jul;39(12):5025-35. II. Yang Y, Hakala H, Mäkelä TP. Nascent RNA analysis reveals the requirement of the mammalian TFIIH kinase Cdk7 for both RNA Pol II and Pol I transcription. Manuscript. III. Helenius K, Yang Y, Alasaari J, Mäkelä TP. Mat1 Inhibits Peroxisome ProliferatorActivated Receptor γ- Mediated Adipocyte Differentiation. Mol Cell Biol. 2009 Jan;29(2):315-23. IV. Yang Y, Roine N, Mäkelä TP. CCRK depletion inhibits glioblastoma cell proliferation in a cilium dependent manner. EMBO Rep. 2013 Aug 1;14(8):741-7. The articles are reproduced with the permission of copyright holders. The author’s contributions: I. Author participated in the experimental design, analysis of results, and preparation of the manuscript. This paper is also included in the theses of Dr. Katja Helenius and Dr. Timofey V. Tselykh. II. Author designed all experiments, performed and analyzed most of the experiments and was the main author of the manuscript. III. Author performed experiments using shRNA targeting Cdk7 and wrote the method section for this; cloned Mat1 cDNA into pBABEpuro retroviral vector. This paper is also included in the thesis of Dr. Katja Helenius. IV. Author designed all experiments, performed and analyzed most of the experiments and was the main author of the manuscript. ii ABBREVATIONS 4sU Ab as ATP Biotin-HPDP bp BrdU CAK CCRK Cdk cDNA CDS ChIP CMV Ct CTD Ctk1 Da DMSO DNA DTT EdU EU FACS FBS FUrd G0 G1 G2 GAPDH GFP GST GTF Hsp70 ICK IF Kin28 M MAK Mat1 4-thiouridine antibody analog-sensitive adenosine triphosphate N-[6-(Biotinamido)hexyl]-3´-(2´-pyridyldithio)propionamide base pair 5-bromo-2'-deoxyuridine cdk activating kinase cell cycle related kinase cyclin-dependent kinase complementary DNA coding sequence for protein chromatin immunoprecipitation cytomegalovirus cycle threshold C-terminal domain of RNA polymerase II Carboxy-Terminal domain Kinase 1 Dalton dimethyl sulfoxide deoxyribonucleic acid dithiothreitol 5-ethynyl-2´-deoxyuridine 5-ethynyl uridine Flow cytometry analysis fetal bovine serum 5-fluorouridine Gap 0 (phase) / quiescence Gap 1 (phase) Gap 2 (phase) glyceraldehyde 3-phosphate dehydrogenase green fluorescent protein glutathionine-S transferase general transcription factor heat shock protein 70 intestinal cell kinase immunofluorescence protein kinase 28 mitosis male germ cell-associated kinase ménage-à-trois 1 iii Mcs MEF mRNA N-terminus NER NT P P-TEFb PAGE PAS PBS PCR PIC Pmh1 Pol polyA PPAR PPRE RNA RNase Rpb rpm RT-qPCR S SDS Ser shRNA siRNA TBP TBS TFII TRC ts TSS XPB/D/F/G mitotic catastrophe suppressor mouse embryonic fibroblast messenger RNA amino-terminus nucleotide excision repair non-targeting phosphorylated positive transcription elongation factor b polyacrylamide gel electrophoresis protein-A sepharose phosphate buffered saline polymerase chain reaction preinitiation complex Pombe Mat1 homolog polymerase polyadenylation peroxisome proliferator-activated receptors PPAR response element ribonucleic acid ribonuclease RNAP II subunit (RNA Polymerase b) rounds per minute real-time quantitative polymerase chain reaction synthesis (phase) sodium dodecyl sulfate serine small hairpin RNA small interfering RNA TATA-binding protein Tris buffered saline transcription factor for RNA polymerase II The RNAi Consortium temperature-sensitive transcription start site xeroderma pigmentosum complementation group B/D/F/G iv ABSTRACT Cyclin-dependent kinases (Cdks) are an evolutionary conserved group of serine/threonine protein kinases involved in critical cellular processes such as cell cycle and transcription. Cdk7 and CCRK (cell cycle related kinase; also known as Cdk20) form a separate branch together in a phylogenetic alignment of Cdks family. This study here has identified distinct cellular functions of these two kinases and does not support overlapping functions as suggested by orthologs in yeast. Cdk7 together with cyclin H and Mat1 forming the kinase subcomplex of TFIIH basal transcription factor complex is proposed to regulate RNA polymerase II (Pol II) mediated mRNA synthesis by phosphorylating the serine-5 (Ser5) residues of Pol II large subunit C-terminal domain (CTD). Investigations in the genetic systems generated here allowing acute depletion of the Cdk7 subcomplex demonstrate that Cdk7 is the mammalian Ser5 phosphorylating kinase and is required for general transcription noted by analysis of newly transcribed RNAs. The analysis also reveals a requirement of Cdk7 for RNA polymerase I-mediated rRNA synthesis. The reduced transcription following Cdk7 disruption is associated with changes on chromatin but not reflected in the steady-state RNA levels due to increased RNA stability. These results also reveal a coupled regulation of transcription and RNA degradation. A tissue-specific function of the Cdk7 subcomplex is identified as a physiological roadblock to adipogenesis by phosphorylating the master transcription factor of adipogenic program-peroxisome proliferator-activated receptor gamma (PPARγ). The observation that the Cdk7 subcomplex is absent from adipose tissues indicates that the socalled basal transcription machinery has very diverse composition in differentiated cells. CCRK is involved in regulating formation of primary cilium, a sensory organelle acting as a signaling hub in the cell. CCRK promotes cell cycle progression by inhibiting ciliogenesis. In glioblastoma cells, reducing the deregulated high level of CCRK or two related substrate kinases of CCRK restores cilia, leading to decreased glioblastoma cell proliferation. Here is identified the first kinase cascade used by tumor cells to disrupt cilia for a growth advantage and offered new therapeutic possibilities. 1 REVIEW OF THE LITERATURE 1. Eukaryotic RNA polymerases Enzymes capable of synthesizing a complementary RNA copy using DNA as template by the polymerization of ribonucleotides are referred as RNA polymerases (Pol). In eukaryotic cells, the task of conveying the genetic information in the nuclear DNA to RNA is shared by RNA Pol I, II and III. The three polymerases were first identified after their chromatographic separation (Roeder and Rutter, 1969, 1970). Subsequent studies on their relative sensitivity to the fungal toxin α-amanitin revealed that they transcribe different sets of genes (Roeder and Rutter, 1970). Pol II produces messenger (m) RNAs coding for proteins and many non-coding RNAs, including small nuclear (sn) RNAs involved in RNA splicing, small nucleolar (sno) RNAs, microRNA (miRNA) precursors, cryptic unstable transcripts (CUTs) (Richard and Manley, 2009) and a novel class long non-coding (lnc) RNAs implicated in regulation of gene expression (Fatica and Bozzoni, 2014). Pol I and III are specialized to transcribe only non-coding RNAs. Pol I exclusively transcribes the multiple copies (~400 in humans) of genes encoding the 18S, 5.8S and 28S ribosomal (r) RNAs. Pol III synthesizes the remaining 5S rRNA and transfer (t) RNAs, the adaptor required in mRNA translation. Other essential Pol III products include 7SL needed for protein insertion into membranes as part of the signal recognition particle and U6 snRNA required for mRNA splicing (White, 2008). The function specificities of the three RNA polymerases are reflected in their subunit compositions (Vannini and Cramer, 2012). RNA Pol II is consisted of 12 highly conserved subunits named Rpb1 to 12 by decreasing order of their molecular mass. Of the 12 subunits, five (Rpb5, Rbp6, Rbp8, Rbp10 and Rbp12) subunits are shared among all three RNA polymerases and four (Rbp1, Rpb2, Rbp3 and Rbp11) have sequencehomologous counterparts in Pol I and III (Thomas and Chiang, 2006). Rpb4, Rbp7, Rbp9 and the carboxy terminal repeat domain (CTD) of Rbp1 are unique in Pol II (Thomas and Chiang, 2006). 2 2. RNA Pol II-mediated synthesis of mRNA 2.1. The unique carboxy terminal repeat domain (CTD) of RNA Pol II The CTD of Rpb1 consists of multiple repeats of an evolutionarily conserved heptapeptide with the consensus sequence Tyr1-Ser2-Pro3-Thr4-Ser5-Pro6-Ser7. Both biochemical and genetic evidences suggest that the CTD serves as a platform in the proximity of the nascent transcript (Meinhart et al., 2005) for the recruitment of factors involved in transcription and co-transcriptional RNA processing (Hsin and Manley, 2012). The number of CTD repeats varies from 26-27 in yeast to 52 in mammals (Allison et al., 1985; Corden et al., 1985) of which eight and 28 repeats required for viability in yeast and human cells respectively (Nonet et al., 1987; West and Corden, 1995). Apparently, the length and diversity in CTD structure increase with the complexity of organism. In most organisms, the majority of CTD heptapeptides match the consensus sequence, while of the 52 CTD repeats in mammals, the first 21 follow the consensus and the last 31 nonconsensus repeats display significant sequence variations at positions 2, 4, 5 and 7 (Corden et al., 1985). It has been suggested that the divergence from the consensus in the C-terminal of the mammalian CTD provides more elaborate regulation of transcription and mRNA processing (Kaneko and Manley, 2005). Many kinds of post-translational modifications can take place at amino acid residues of CTD of Rpb1, among which phosphorylations of serine 2 and 5 (Ser2P and Ser5P) are most extensively studied. The phosphorylation of Ser7 (Ser7P) and other covalent modifications identified more recently appear to associate with gene-specific functions (Egloff et al., 2012). Ser2 and Ser5 of CTD are subjected to dynamic phosphorylation and dephosphorylation creating a characteristic CTD phosphorylation pattern correlated with the transcription cycle. The dynamics appears to be essential as the substitution of Ser2 and Ser5 with alanine or an acid mimic (glutamate) in S. cerevisiae is lethal (West and Corden, 1995). Multiple functions for Ser2P and Ser5P in transcription have been identified and are discussed in detail in the section 2.3. Although it has been shown that the CTD phosphorylation does not influence the inherent catalytic activity of Pol II (Serizawa et al., 1993), multiple lines of evidence indicate that the phosphorylation of the 3 CTD is critical for various steps of mRNA synthesis and processing in the context of chromatin (to see the section 2.3). 2.2. Role of chromatin structure in transcription In eukaryotes, DNA undergoes a series of compactions in association with specific proteins to form chromatin (Cockerill, 2011). The first level of compaction involves 147 base pairs of DNA wrapped around two copies of the core histones H2A, H2B, H3 and H4 to form a structure called nucleosome. This produces a 10 nm fiber with a beads-on-astring structure. The N-terminus of histones protruding from the nucleosome is a target for a wide array of modifications including acetylation, methylation and phosphorylation. The modifications are applied and removed in a highly specific and sometimes coordinated manner to generate a “histone code” which can be recognized by chromatinassociated factors. Further compaction involves histone modifications and a linker histone H1 occupying the DNA between nucleosomes, producing a 30 nm fiber with a helical structure of nucleosomes. The 30 nm fiber is further compacted by forming loops tightly packed in heterochromatin and ultimately the mitotic chromosome during cell division. The compacted chromatin structure presents an inherent obstacle to transcription by preventing DNA accessibility. Localized chromatin decondensation and nucleosome displacement are required to create a chromatin environment allowing the passage of polymerases (Cockerill, 2011). The key regulatory DNA sequences near the promoter and transcriptional start sites are often depleted of nucleosomes (Yuan et al., 2005). The elongating Pol II must break the nucleosome barrier in the coding region to gain access to DNA and transcribe genes efficiently. This involves recruiting histone modifiers and chromatin remodelers to slide nucleosomes and evict histones (or deposit histone variants), as well as histone chaperons to facilitate the disassembly and reassembly of nucleosomes (Petesch and Lis, 2012). The CTD modulated by phosphorylation provides a means to recruit these factors to regulate transcription as discussed in detail in the section 2.3. 2.3. The RNA polymerase II transcription cycle and CTD phosphorylation Pol II transcription plays a central role in gene expression and is extensively regulated at 4 each step from initiation, promoter escape, elongation to termination. The Ser2 and Ser5 phosphorylation of CTD is involved in all transcription steps and coordinates these events with co-transcriptional RNA processing (capping, splicing and polyadenylation). Pol II-mediated transcription begins with formation of the preinitiation complex (PIC) at the gene promoters, containing hypophosphorylated RNA Pol II and a set of general transcription factors (GTFs) recruited in an ordered sequence (TFIID, TFIIA, TFIIB, TFIIF, TFIIE and TFIIH) (Thomas and Chiang, 2006). The classic model for PIC assembly starts with the recruitment of the TATA-binding protein (TBP) contained in TFIID complex to the TATA box core promoter element (Buratowski et al., 1989). The formation of PIC is regulated by gene-specific activators and co-activators. Of the limited number of coactivators known to directly target Pol II, the multiprotein complex Mediator is potentially the most crucial regulator due to its widespread and multifaceted functions across the transcriptome (Malik and Roeder, 2010). In the PIC, Pol II is anchored to promoter by the DNA-GTF interactions and cannot initiate transcription until the template DNA strands are separated. The XPB subunit of TFIIH is required for ATPdependent unwinding of about ten base pairs (bp) of promoter DNA, forming the initial transcription bubble (Kim et al., 2000). Pol II then locates the transcription start site (TSS) and undergoes promoter escape. The multiple interactions between Pol II and other PIC components are thought to be disrupted by Ser5P, permitting Pol II to escape from the promoter (Akoulitchev et al., 1995; Jiang et al., 1996; Sogaard and Svejstrup, 2007). A more clearly defined role of Ser5P is recruiting and activating the capping machinery to place the 7-methyl-guanosine cap on the 5’ end of the nascent RNAs (Cho et al., 1997; McCracken et al., 1997). In S. pombe, the capping enzyme interacts with and recruits the Ser2 phosphorylating kinase to chromatin likely to ensure that only properly capped transcripts are elongated (Guiguen et al., 2007; Viladevall et al., 2009), given the importance of Ser2P in productive elongation (see below). The lethality caused by substituting Ser5 to alanine can be circumvented by covalent tethering of mRNA-capping enzymes to the CTD in S. pombe (Schwer and Shuman, 2011), suggesting that recruitment of the capping machinery is the essential function of Ser5P. Ser5P is also required for interaction with the yeast Nrd1, a factor 5 involved in the 3’ end formation and early termination of non-polyadenylated transcripts (Gudipati et al., 2008; Vasiljeva et al., 2008). The histone methyltransferase Set1, which trimethylates histone H3 lysine 4 (H3K4trime) indicating active transcription, interacts with the early elongating Pol II through Ser5 phosphorylated CTD in S. cerevisiae (Ng et al., 2003). Ser5P also stimulates co-transcriptional recruitment of histone deacetylase in S. cerevisiae to prevent excessive nucleosome eviction (Govind et al., 2010). The level of Ser5P peaks early in the transcription cycle and remains constant or decreases as the elongating Pol II progresses to the 3’ end of the gene. In metazoans, the transition to productive elongation frequently involves promoter-proximal pausing of the Ser5 phosphorylated Pol II at many developmentally and environmentally responsive genes (Adelman and Lis, 2012) and is often regulated to ensure synchronous gene expression (Boettiger and Levine, 2009). The paused Pol II remains stably associated with capped nascent RNA (ca. 50 nt) and is competent to resume elongation after being released. The Ser2 phosphorylating kinase Cdk9 has been demonstrated to play a major role in overcoming this early elongation block and the conversion into a productive elongation (Peterlin and Price, 2006). Subsequent to the Ser5P enrichment at the promoter region, CTD Ser2P increases towards the 3’ end and plays a major role in recruiting factors associated with productive elongation and factors required for splicing, polyadenylation, cleavage and termination. The Ser2 and Ser5 double phosphorylated CTD is directly recognized by the splicing factor U2AF65, which in turn recruits the splicing factor Prp19 to nascent RNA for cotranscriptional splicing (David et al., 2011). Nucleosome reassembly during elongation is coordinated by several histone modifications, of which trimethylation of histone H3 at lysine 36 (H3K36trime) is mediated by Set2 histone methyltransferase that is recruited to chromatin also by the Ser2P and Ser5P double mark in yeast (Kizer et al., 2005; Vojnic et al., 2006). The murine Spt6 elongation factor that also functions as histone H3 chaperone in nucleosomes reassembly interacts directly with Ser2 phosphorylated CTD of Pol II (Yoh et al., 2007). The interaction of Spt6 with Ser2P facilitates the recruitment of the nuclear RNA export factor REF1/Aly in mammalian cells (Yoh et al., 2007). Ser2P is also required for maintaining H2B monoubiquitination and correct 3’-end processing of 6 replication-dependent histone mRNAs (Pirngruber et al., 2009). Transcription termination of mRNA-encoding genes is coupled to 3’-end processing of the transcripts. In mammals, the 3’ end of nascent mRNA is first cleaved endonucleolytically at the polyadenylation (polyA) site between two specific sequence elements (a conserved hexanucleotide AAUAAA and a less conserved U- or GU-rich region) followed by the addition of a poly(A) tail consisting of multiple adenosine monophosphates to the free 3’ end (Proudfoot, 2011). The uncapped residual RNA associated with Pol II that transcribes beyond the polyA site is rapidly degraded by the exonuclease, leading to Pol II release and transcription termination (Proudfoot, 2011). The yeast Pcf11 is an essential polyadenylation factor that preferentially binds to Ser2 phosphorylated CTD, which may help the elongation Pol II scan for the specific elements (Licatalosi et al., 2002). Ser2P has also been suggested to help recruitment of the yeast exonuclease Rat1 (Kim et al., 2004b). Interestingly, Ser2P in contrast to Ser5P antagonizes the recruitment of Nrd1 and Nrd1-dependent termination (Gudipati et al., 2008). Higher Ser2P compared to Ser5P in Pol II later in transcription thus favors the canonical mRNA 3’-end processing, suggesting that the phosphorylation status of CTD dictates the choice of termination pathway (Gudipati et al., 2008). Following the release of Pol II from the template, the CTD is dephosphorylated enabling another round of transcription (Cho et al., 1999). In summary, the changes in Ser2P and Ser5P of Pol II CTD couple each phase of the transcription cycle to the corresponding RNA processing events through dynamic interactions with distinct sets of factors. 2.4. Mutual regulation of transcription and decay As discussed in the previous section, transcription and the various precursor mRNA (premRNA) processing events are tightly coupled to ensure production of mature mRNAs. A failure at any stage may cause RNA retention in the nucleus and trigger its degradation. For example, the quality of the 5’ end cap methylation is monitored by the Rat1/Rai1 complex and the pre-mRNAs with improperly methylated caps are degraded before exported to the cytoplasm (Jiao et al., 2010). The degradation of splicing-defective premRNAs in the nucleus involves a multi-protein complex known as the exosome 7 containing 3’ to 5’ exonuclease activity (Bousquet-Antonelli et al., 2000). Aberrant mRNAs including splicing-defective pre-mRNAs that escape to the cytoplasm are degraded by pathways of mRNA surveillance or nonsense-mediated decay (NMD) (Schoenberg and Maquat, 2012). In the case of functional mRNAs, the degradation in the cytoplasm involves prior deadenylation of the mRNAs. This occurs after the mRNA has been translated by ribosomes to produce functional proteins. Following removal of the poly(A) tail by a deadenylase activity, the mRNA can be degraded in a 3’ to 5’ direction by the exosome (Anderson and Parker, 1998) with the remaining cap structure being hydrolyzed by the scavenger-decapping enzyme (Liu et al., 2002a). Alternatively, the deadenylated mRNA undergoes decapping first allowing the mRNA susceptible to decay by the 5’ to 3’ exoribonuclease Xrn1 (Hsu and Stevens, 1993; Muhlrad et al., 1994). These two pathways are not mutually exclusive and the respective contribution remains unknown (Garneau et al., 2007). Maintaining appropriate levels of mRNAs is essential for any living cells. The steadystate levels of mRNAs in a cell are determined by the balance between synthesis and degradation. Recently several reports have demonstrated that transcription and mRNA decay are coupled in yeast (Dahan and Choder, 2013; Haimovich et al., 2013; Sun et al., 2013; Sun et al., 2012) through at least Pol II subunits Rpb4/7 (Goler-Baron et al., 2008), a transcription activator Rap1 (Bregman et al., 2011) and the decay factor Xrn1 (Haimovich et al., 2013; Sun et al., 2013). The two Pol II subunits Rpb4 and Rpb7 strongly interact with each other forming a heterodimer, a substructure distinct from the “core Pol II” consisted of the other ten subunits and required for promoter-dependent transcription in vitro (Edwards et al., 1991). Rpb4 and Rpb7 are present in excess over the core subunits and shuttle between the nucleus and the cytoplasm, suggesting that they might function separately from Pol II (Selitrennik et al., 2006). Indeed Rpb4 and Rpb7 have been implicated in mRNA export (Farago et al., 2003), translation (Harel-Sharvit et al., 2010) and degradation in S. cerevisiae (Goler-Baron et al., 2008; Lotan et al., 2005; Lotan et al., 2007). Remarkably, the yeast Rpb4/7 dimer interacts with nascent RNAs emerging from Pol II during 8 transcription, dissociates from Pol II together with the transcripts and remains bound throughout the mRNA lifecycle (Dahan and Choder, 2013). Rpb4/7 directly functions in mRNA deadenylation and regulates the two mRNA decay pathways (Lotan et al., 2005; Lotan et al., 2007). Interestingly, in strains with mutant forms of Pol II defective in recruiting Rpb4/7, the interaction of Rpb4/7 with transcripts is compromised as well as deadenylation and mRNA decay, suggesting the mRNA interaction of Rpb4/7 occurs only in the context of Pol II and is required for Rpb4/7-mediated stimulation of mRNA decay (Goler-Baron et al., 2008). These pioneering findings reveal that the transcriptional machinery can remotely control the mRNA stability in cytoplasm through imprinting the transcript with factors that escort it to the cytoplasm and regulate its degradation. Interestingly, a yeast strain harboring defective Pol II mutant with slower transcription rate is also slow in mRNA degradation, resulting in a buffering of mRNA levels (Sun et al., 2012). A similar phenomenon has been observed following depletion of Rap1 (Bregman et al., 2011). The underlying mechanisms however are unknown and it would be interesting to examine the mRNA imprinting status in this mutant strain. Vice versa, a decrease in decay rate caused by deletion of the decay factors is compensated by a decrease in transcription rate to maintain the steady-state mRNA levels in S. cerevisiae (Haimovich et al., 2013; Sun et al., 2013). Xrn1 has been identified as the key regulator of such coupling although the mechanisms by which Xrn1 regulates transcription remain controversial (Haimovich et al., 2013; Sun et al., 2013). Interestingly, this coupling between transcription and cytoplasmic decay might be achieved through gene promoters. Rpb4/7 is present during initiation at promoters (Jasiak et al., 2008), components of the 5’ to 3’ Xrn1 decay pathway have been suggested to enrich at promoter regions (Haimovich et al., 2013). Interestingly, two studies in yeast demonstrate that modulation of promoter sequences influences the stability of the transcribed mRNAs (Bregman et al., 2011; Trcek et al., 2011). A similar promoterregulated mRNA turnover might also exist in mammalian cells as suggested by an early study on a single human gene (β-globin) (Enssle et al., 1993). It is not known yet whether promoter-dependent regulation of mRNA stability could be a common strategy. The 9 molecular basis for how promoters determine the fate of mRNAs also remains unknown. Importantly, it is unknown whether the coordination between transcription and mRNA degradation also exist in organisms other than S. cerevisiae. The coupling between transcription and decay might play an important role in response to environmenal stimuli as many environmentally responsive genes are subject to transcription induction accompanied by increase in decay (Dahan and Choder, 2013). The bilateral dialog between transcription and mRNA decay might represent another layer of regulation of gene expression to maintain the desired steady-state levels of mRNAs or shape the kinetics of achieving new mRNA levels in response to environmental stimuli. 2.5. Transcriptional cyclin-dependent kinases (Cdks) phosphorylate Pol II CTD Members of the Cdk family are the major CTD phosphorylating kinases. As their names indicate, Cdks are supposed to be kinases that depend on their associated regulatory subunit, the cyclins, for proper function. Current nomenclature for Cdks does not follow the strict ‘cyclin-dependent’ rule, instead is based on sufficient sequence similarities (Malumbres et al., 2009). Twenty CDKs have been identified to be encoded by the human genome (Figure 1), of which Cdk7, Cdk8, Cdk19, Cdk9 and Cdk12 have been suggested to phosphorylate CTD of Pol II. 10 Figure 1. A rooted phylogenetic tree with branch length of human CDKs based on the amino acid sequences. As shown, CDK7 and CCRK, CDK8 and CDK19, CDK9 together with CDK12 and CDK13 cluster in separate branches. The amino acid sequences of human CDK1 (P06493, isoform 1), CDK2 (P24941, isoform 1), CDK3 (Q00526), CDK4 (P11802), CDK5 (Q00535, isoform 1), CDK6 (Q00534), CDK7 (P50613) CDK8 (P49336, isoform 1), CDK9 (P50750, isoform 1), CDK10 (Q15131, isoform 1), CDK11A (Q9UQ88, isoform SV6), CDK11B (P21127, isoform SV9), CDK12 (Q9NYV4, isoform 1), CDK13 (Q14004, isoform 1), CDK14 (Q94921, isoform 1), CDK15 (Q96Q40, isoform 1), CDK16 (Q00536, isoform 1), CDK17 (Q00537, isoform 1), CDK18 (Q07002, isoform 1), CDK19 (Q9BWU1, isoform 1) and CCRK (Q8IZL9, isoform 1) were used in CLUSTALW Multiple Sequence Alignment (http://www.genome.jp/tools/clustalw/; March, 2014) with default settings. Mammalian Cdk7 was identified as a CTD kinase also possessing in vitro Cdk-activating kinase (CAK) activity phosphorylating the T-loop threonine of other Cdks involved in cell cycle regulation (Roy et al., 1994; Serizawa et al., 1995; Shiekhattar et al., 1995). However, the homologue of Cdk7 in S. cerevisiae Kin28 lacks CAK activity (Cismowski et al., 1995) and the CAK activity is instead mediated by a structurally unrelated kinase termed Cak1 (Espinoza et al., 1996; Espinoza et al., 1998; Kaldis et al., 1996). The discovery of the alternative CAK in yeast prompted the re-evaluation of Cdk7 as the in vivo CAK in metazoans. The cell cycle-related kinase (CCRK) or Cdk20 is regarded as a candidate because it is the most closely related kinase to Cdk7 in a phylogenetic alignment of Cdk family (Guo and Stiller, 2004; Malumbres et al., 2009). CCRK has been implicated in cell cycle regulation but whether it regulates the cell cycle as a CAK remains controversial (Feng et al., 2011). Notably, no kinase activity of CCRK has been detected towards CTD in vitro (Wohlbold et al., 2006 and our unpublished results). The 11 role of Cdk7/Kin28 in transcription as a CTD phosphorylating kinase is discussed in detail in the section 2.6. Cdk8-cyclin C is part of the Mediator complex recruited to promoters and can phosphorylate CTD Ser5 in vitro (Akoulitchev et al., 2000). Cdk19 (previously called Cdk11) was discovered by mass spectrometry as a Mediator-associated kinase (Sato et al., 2004) that is conserved only in vertebrates (Tsutsui et al., 2008). Cdk19 shares high degree of sequence identity also the counterpart cyclin C with Cdk8 and in turn has been suggested to function redundantly, collaboratively or competitively with Cdk8 depending on contexts (Galbraith et al., 2013; Tsutsui et al., 2011). A Ser5-specific kinase activity of Cdk19 similar to Cdk8 has been shown in vitro (Tsutsui et al., 2011). However, the in vivo relevance of the Ser5-phosphorylating capacity of Cdk8 and Cdk19 remains to be defined. Cdk9 is the major Ser2 kinase in metazoans as the catalytic subunit of the positive transcription elongation factor b (P-TEFb) that controls the elongation phase (Peterlin and Price, 2006). S. cerevisiae has two Ser2 CTD kinases, Bur1 and Ctk1 (Wood and Shilatifard, 2006). There are two orthologs of yeast Ctk1 in humans, Cdk12 and Cdk13 sharing 89% sequence identity; so far only Cdk12 has been clearly demonstrated to be an elongating CTD kinase (Bartkowiak et al., 2010). The transcriptional Cdks are recruited to chromatin at different steps of transcription, and with the help of CTD phosphatases generate the characteristic CTD phosphorylation pattern correlating with the position of Pol II along genes. In addition, these Cdks are able to modulate each other’s activity. Mammalian Cdk8 has been shown to phosphorylate the cyclin partner of Cdk7 (cyclin H) resulting in inhibition of the CTD kinase activity of Cdk7 (Akoulitchev et al., 2000). Recently, Cdk7 has been reported to phosphorylate the T-loop threonine of Cdk9 and this phosphorylation seems to be required for the activity of Cdk9 in phosphorylating Ser2 of CTD in mammalian cells (Larochelle et al., 2012). The interplay between the transcriptional Cdks suggests a CDK network controlling CTD phosphorylations and thus regulating Pol II-mediated transcription. 12 2.6. Cdk7-cyclin H-Mat1 as the kinase subcomplex of TFIIH in general transcription Cdk7 forms a complex with cyclin H and a RING finger protein ménage-à-trois 1 (MAT1) as the kinase submodule of TFIIH. Mammalian TFIIH consists of ten subunits that make up the core (XPB, XPD, p8/TTDA, p34, p44, p52, p62) and the CAK subcomplex composed of Cdk7-cyclinH-Mat1. In addition to transcription as mentioned previously, TFIIH also functions in nucleotide excision repair (NER) during which the Cdk7 kinase subcomplex is released from the core and back together with the resumption of transcription after the complete removal of DNA lesions (Coin et al., 2008). These observations imply that Cdk7 is not required for the TFIIH core structure or its function in NER but is essential as a coordinator between transcription and NER, yet the mechanism of how the dynamic organization of TFIIH complex is achieved in regard to the physiologic changes is unknown. Cdk7-cyclinH-Mat1 is recruited to the preinitiation complex (PIC) as a part of TFIIH and is thought to regulate general transcription through phosphorylating Ser5 of Pol II CTD (Serizawa et al., 1995; Shiekhattar et al., 1995). Recent studies have identified Cdk7/Kin28 phosphorylating Ser7 (Akhtar et al., 2009; Glover-Cutter et al., 2009; Kim et al., 2009), which is required for the transcription of snRNA genes but not for proteinencoding genes (Chapman et al., 2007; Egloff et al., 2007). There have been conflicting observations concerning that Cdk7 is the CTD Ser5 kinase in vivo and the resulted Ser5P is essential for global mRNA transcription and processing. At the non-permissive temperature, thermosensitive mutant of Cdk7/Kin28 in S. cerevisiae (Valay et al., 1995) and C. elegans (Wallenfang and Seydoux, 2002) displayed a dramatic decrease in Ser5P of Pol II CTD as well as transcription. In contrast, chemical inhibiting ATP-analogue sensitive mutant of Kin28 in S. cerevisiae showed decreased Ser5P and 5’capping of transcripts, but no major effects on mRNA abundance (Akhtar et al., 2009; Kanin et al., 2007), suggesting that Cdk7/Kin28 activity or CTD Ser5P is not needed in transcription per se but rather required for mRNA maturation. However, later using the same methods, another group reported that inhibition of Kin28 activity does cause global impairment of gene expression and the reduced mRNA level is correlated with the 13 reduction in Ser5P (Hong et al., 2009). In S. pombe, thermal or chemical inactivation of the Mcs6/Mcs2/Pmh1 complex (orthologous to Cdk7/cyclin H/Mat1) caused genespecific rather than global defects in transcription associated with decreased Ser5P (Lee et al., 2005; Viladevall et al., 2009). Analysis of a Drosophila Cdk7 temperature-sensitive mutant indicated a requirement for full induction of the heat shock genes but not all the genes (Schwartz et al., 2003). In mice, Mat1-null cells exhibit reduced Ser5 phosphorylation (Rossi et al., 2001) and deletion of Mat1 in murine Schwann cell precursors does not inhibit the transcription program of myelination (Korsisaari et al., 2002). On the contrary, chemical inhibition of Cdk7 in a human cell line or depletion of Cdk7 in mice does not decrease the cellular level of Ser5P (Ganuza et al., 2012; GloverCutter et al., 2009; Larochelle et al., 2012). In summary, the extent of Cdk7 contributing to global mRNA expression in vivo remains controversial and its role as the major Ser5 phosphorylation kinase in mammalian cells is challenged by the recent studies in mammalian systems (Ganuza et al., 2012; Glover-Cutter et al., 2009; Larochelle et al., 2012). 2.7. Cdk7-cyclin H-Mat1 in cell type-specific transcription The stimulation of transcription program driving differentiation to generate different cell types is achieved by binding of gene-specific transcription factors to the regulator DNA sequences in response to developmental and environmental signals. The binding stimulates the general transcription machinery to produce cell type-specific transcripts mediating the phenotype. The role of these specific transcription factors is instructive as modulation of their expression or activity can often determine cell fate. By contrast, the general transcription machinery has largely been relegated as a supporter that only conveys the instructions from the specific transcription factors. TFIIH-associated Cdk7 can phosphorylate gene-specific transcription factors including peroxisome proliferator-activated receptor γ (PPARγ) at serine 112 (S112) in vitro (Compe et al., 2005). PPARγ is the master transcription factor of the adipogenic transcription program, as its expression is necessary and sufficient to drive adipocyte differentiation (Rosen et al., 1999; Tontonoz et al., 1994). The S112 in the N-terminal 14 activation domain of PPARγ is an inhibitory site originally identified as mitogenactivated protein kinase site (Adams et al., 1997; Camp and Tafuri, 1997; Hu et al., 1996). Phosphorylation of PPARγ S112 represses the activity PPARγ and inhibits the transcriptional activation of PPARγ target genes (Shao et al., 1998). Furthermore, mutated S112 to non-phosphorylatable alanine promotes adipocyte differentiation (Hu et al., 1996; Rangwala et al., 2003). The activity of PPARγ is linked to Cdk7 in vivo by the observation that expression of PPARγ target genes are deregulated in adipose tissue of mice with mutations in the XPD subunit of TFIIH (Compe et al., 2005). It is of great interest to study the potential role of Cdk7 in adipogenic program mediated by PPARγ as this may reveal possible contributions of the basal transcription machinery to orchestrate cell type-specific gene expression. 3. RNA polymerase I-mediated synthesis of rRNA In vertebrates, rRNA genes (rDNA) are transcribed by Pol I in the nucleolus into a precursor RNA (45S or 47S pre-rRNA) processed into mature 18S, 5.8S and 28S rRNAs that are essential structural and catalytic components of ribosomes, the core protein synthetic machinery. Ribosome biogenesis is a highly conserved and energy-consuming process in the nucleolus. Given the metabolic burden and its direct impact on protein synthetic capacity, Pol I-mediated transcription of rRNA is subjected to a tight regulation to ensuring that rRNA production is closely coupled with cellular states (Goodfellow and Zomerdijk, 2012). 3.1. The structure of rRNA genes The rDNA exists as tandem repeats in all eukaryotes and the composition of each repeat is conserved despite varieties in the number of repeats and the size of the gene. In yeast, there are about 200 repeats in a single locus per haploid genome whereas in humans about 360 copies of ribosomal genes are located in five chromosomal loci termed nucleolar organizer regions (NORs) (Birch and Zomerdijk, 2008). In mice about 200 rDNA repeats are distributed among five chromosomes (Grozdanov et al., 2003). Only a faction of these repeats is actively transcribed and distinguished from the inactive repeats by distinct chromatin structures, epigenetic marks and sub-nucleolar localization 15 (Goodfellow and Zomerdijk, 2012). The inactive rDNA is associated with a tightly packed heterochromatin state characterized by DNA methylation and repressive histone modifications, and tethered to the nucleolar periphery (Goodfellow and Zomerdijk, 2012). The active rDNA is in a more open chromatin structure with hypomethylated DNA, acetylated histones and an enrichment of the Pol I activator upstream binding factor (UBF), and transcribed at the nucleolar fibrillar center (Fakan and Hernandez-Verdun, 1986; Scheer and Rose, 1984). A rDNA repeat is about 43 kbp in humans and 45 kbp in mice containing regulatory elements within an intergenic spacer (IGS) of ~30 kb and a single transcribed region of ~13 kb encoding pre-rRNA (McStay and Grummt, 2008). Notably, the multiple rDNA repeats are not identical copies; several rDNA variants exist and can be differentially expressed and regulated (Tseng et al., 2008). Regulatory elements include promoters, spacer promoters, repetitive enhancers and terminators. The rDNA promoter is consisted of a core promoter element adjacent to the TSS and an upstream control element (UCE) ~ 100 bp further upstream, both of which are important for directing accurate and efficient initiation of pre-rRNA synthesis. The promoter is immediately preceded by a transcriptional terminator sequence (T0) required for efficient initiation (Grummt et al., 1986). Upstream of T0 are multiple copies of a repetitive sequence called enhancer repeats that can stimulate transcription (Pikaard et al., 1990). Upstream of enhancer repeats, a functional element called spacer promoter has been identified in several species and suggested to stimulate transcription form the sequencerelated rDNA promoter in mice (Paalman et al., 1995), although the transcripts produced from this promoter are implicated in epigenetically silencing of a subset of rDNA arrays (Mayer et al., 2006). 3.2. Pol I transcription processes on rDNA Transcription of rDNA by Pol I is initiated by assembly of a PIC on the promoter, including binding of UBF and the promoter selectivity factor 1 (SL1) in humans or TIFIB termed in mice (Clos et al., 1986; Learned et al., 1986). UBF can activate rDNA transcription by recruiting Pol I to the rDNA promoter, stabilizing binding of SL1/TIF-IB 16 and displacing histone H1 (Bell et al., 1988; Kuhn and Grummt, 1992). More recently, UBF has been suggested to regulate Pol I promoter escape (Panov et al., 2006) and transcription elongation (Stefanovsky et al., 2006). SL1/TIF-IB is a ~300 kDa protein complex containing TBP and at least four Pol I-specific TBP-associated factors (TAFs) mediating specific interactions between the rDNA promoter and Pol I. An additional TAF, TAF12 originally described as Pol II-specific has been identified in SL1/TIF-IB complex involved in Pol I transcription (Denissov et al., 2007). These TAFs interact with UBF and recruit Pol I by binding to TIF-IA, the mammalian homolog of yeast Rrn3 associated with the initiation-competent subpopulation of Pol I (Pol I β) (McStay and Grummt, 2008). Numerous proteins including kinases, chromatin modifiers, nuclear actin and myosin as well as proteins involved in DNA repair are associated with Pol I β, suggesting that Pol I is recruited to the rDNA promoter as a massive multiprotein complex that acts as a scaffold to coordinate rRNA synthesis and maturation as well as chromatin modification and DNA repair (Drygin et al., 2010). Interestingly, some of these proteins are shared between the Pol I and Pol II transcription machinery, including TFIIH (Assfalg et al., 2012; Bradsher et al., 2002; Hoogstraten et al., 2002; Iben et al., 2002). Pol I has intrinsic RNA hydrolysis activity and elongation rate-enhancing subunits to promote transcript cleavage in an arrested complex during transcription elongation (Kuhn et al., 2007), whereas Pol II required TFIIS to clear arrests. Pol I also contains subunits that functionally resemble TFIIF in regulating elongation (Albert et al., 2011; Beckouet et al., 2008; Kuhn et al., 2007). For transcription termination, a major terminator sequence located downstream from the pre-rRNA coding sequence is recognized by the DNA-binding factor Transcription Termination Factor I (TTF-I) in mammals (Evers et al., 1995). The binding of TTF-I to the terminator sequence causes Pol I pausing and the paused Pol I is dissociated by release factor (Jansa and Grummt, 1999; Jansa et al., 1998; Mason et al., 1997). However a more complicated control of Pol I termination has been suggested by recent studies in yeast (Nemeth et al., 2013). 17 3.3. Involvement of modifiers of Pol II CTD in rDNA transcription The pre-rRNA transcribed by Pol I is subject to extensive co-transcriptional modification and a cascade of endo/exonucleolytic cleavage and degradation steps that are tightly controlled to produce mature 18S, 5.8S and 28S rRNA for ribosome biogenesis. Interestingly, a recent study reported that knockdown of Cdk9 inhibits rRNA processing and this inhibition was proposed to be secondary to a defect in Pol-II mediated transcription of a snoRNA that is required for rRNA processing (Burger et al., 2013). It appears that the enzymes catalyzing phosphorylation and dephosphorylation of Pol II CTD are implicated in Pol I transcription. The dual function of these proteins might provide a means to coordinate the control of mRNA and rRNA synthesis in eukaryotic cells. The Cdk7 homolog Kin28 is required for rRNA synthesis in S. cerevisiae (Iben et al., 2002). The mammalian TFIIH including the Cdk7 kinase submodule can be isolated in a complex with Pol I and TIF-IB (Bradsher et al., 2002; Hoogstraten et al., 2002; Yuan et al., 2007). Photobleaching experiments demonstrated that the core TFIIH subunit XPB resides 25 s for average time in the nucleolus whereas 2-10 s at a Pol II promoter, suggesting that the individual TFIIH complex has a longer engagement in the Pol I transcription machinery than in the Pol II machinery (Hoogstraten et al., 2002). Consistently, the core and the Cdk7 subcomplex of TFIIH were found to reside at geneinternal sequences and leave the rDNA promoter with Pol I after transcription initiation (Assfalg et al., 2012). Mutations in the core subunits XPB and XPD found in Cockayne syndrome disrupt the rDNA association of TFIIH, but do not affect PIC or promoter escape of Pol I in vitro, suggesting that TFIIH might function in rRNA transcription at elongation stage rather than initiation (Assfalg et al., 2012). The contributions of the Cdk7 kinase submodule to rRNA transcription in mammalian cells remain to be established. In addition to Kin28, the Ctk1 kinase in yeast has also been shown to interact with Pol I and function in rRNA transcription (Bouchoux et al., 2004). A CTD phosphatase Fcp1 that functions in the Pol II recycling is also part of the Pol I transcription machinery and is required for efficient rRNA synthesis (Fath et al., 2004). It is tempting to speculate that 18 this phosphatase may antagonize the activity of CTD kinases to produce a similar phosphorylation cycle in the transcription mediated by Pol I as that mediated by Pol II. The CTD repeats are unique to Pol II and the substrates for these kinases and the phosphatase Fcp1 in Pol I transcription remain unknown. 4. Regulation of ciliogenesis and links to cell cycle control 4.1. The structure of primary cilia Eukaryotic cilia and flagella are microtubule-based organelles covered by a layer of specialized membrane protruding from cell surfaces. Cilia are conventionally classified as primary (immotile) and motile cilia. Primary cilia, motile cilia and flagella are remarkably conserved in their structures from unicellular eukaryotes to mammals, but their number, length and functions vary from one cell type to another. Flagellar and the typical motile cilia have the identical axonemal structures for their locomotive ability: a “9+2” microtubule arrangement (nine doublet microtubules arranged in a circle surrounding two central singlet microtubules), radial spokes and dynein arms attached to the microtubule doublets viewed in cross section (Figure 2). Flagella use an undulating motion to propel cells forward while motile cilia beat back and forth to sweep materials across cell surfaces. Flagella are found on unicellular eukaryotes and sperms. The mammalian spermatozoon has a single flagellum while the green alga Chlamydomonas has two flagella and the unicellular protozoan Tetrahymena is covered with a few thousand. In mammals, ciliated cells with up to 200 motile cilia are found in the respiratory tract and oviduct required for mucus clearance and egg movement respectively (Lodish et al., 2000). Primary cilia usually have a “9+0” axonemal structure consisting of a ring of nine doublets without the central pair of microtubules. The radial spokes and dynein arms are also not present in primary cilia (Figure 2) (Ishikawa and Marshall, 2011). Primary cilia are non-motile and were considered an evolutionary remnant until the findings within the past decades linked them to cell signaling transduction (Huangfu and Anderson, 2005) and various human diseases collectively called “ciliopathies” (Badano et al., 2006). These 19 findings have revived the interest in studying this overlooked organelle. Since then evidences have piled up highlighting primary cilia as an essential sensor for mechanical or chemical signals and as a signaling platform for several important signaling pathways. A single primary cilium is present on the apical surface of most of mammalian cells, including epithelial, endothelia, bone and muscle cells as well as neurons (http://www.bowserlab.org/primarycilia/cilialist.html; March, 2014). Although primary cilia do not provide cell mobility, they can be bent in response to flow or fluid shear stress. For example, the bending of primary cilia in renal epithelial cells induces calcium flux (Praetorius and Spring, 2001). Notably, recent findings also identify exceptional motile “9+0” cilia (e.g. nodal cilia). The motile “9+0” cilia at the embryonic node generate a leftward flow and act with the immotile cilia involved in sensation of nodal flow together to establish the left-right asymmetry (McGrath et al., 2003). However, it is still controversial whether the nodal cilia can also sense nodal flow. There are also studies suggesting a sensory function of other motile “9+2” cilia, thus the functional distinction between the motile and primary cilia is not strict (Bloodgood, 2010). Primary cilia are anchored to cells through a centriole-derived structure called “basal body”. During the conversion to basal body, the mother centriole generates specialized appendages at its distal end that allow it to interact with the membrane and nucleate the axoneme. The ciliary membrane despite being continuous with the plasma membrane maintains a distinct lipid composition and is enriched in proteins implicated in signaling pathways (Garcia-Gonzalo and Reiter, 2012). The membrane invagination next to the ciliary base is called “ciliary pocket”. The basal body is anchored to the ciliary membrane at the curved base of the ciliary pocket by transition fibers - a pinwheel-like structure identified on transmission electron microscope cross sections (Anderson, 1974). Other structures, such as basal feet and ciliary rootlet, originate from the proximal ends of basal body and interact with cytosolic cytoskeleton, providing structural support for cilia (Yang et al., 2005). 20 IFT complex B IFT particle remodeling & microtubule turnover ‘Core’ ‘Peripheral’ IFT52 IFT25 IFT20 IFT70 IFT46 IFT54 IFT74 IFT57 IFT81 IFT80 IFT88 IFT172 IFT22 (Rabl5) IFT27 (Rabl4) GTPase TRP BBS4 BBS8 GTPase BBS3 PH PH BBS5 KAP KIF3A KIF3B Ciliary membrane Cross section of middle segment (primary cilia) IFT-dynein motor heavy chain (DYNC2H1/DNCHC2) intermediate chain (WDR34) light intermediate chain (DYNC2LI1) light chain (DYNLL1) Anterograde β-propeller BBS1 BBS2 BBS7 BBS9 A tubule Homodimeric Kinesin-2 KIF17(OSM-3) Heterotrimeric Kinesin-2 Retrograde BBSome Cross section of distal segment (primary cilia) Cross section of middle segment (motile cilia and flagella) Nexin Outer dynein arm B tubule IFT complex A ‘Core’ IFT122 IFT140 IFT144 ‘Peripheral’ IFT43 IFT121 IFT139 Y-link Radial spoke Central pair Inner dynein arm Plasma membrane Ciliary pocket Transition zone Transition fibers Basal body C tubule basal feet ciliary rootlet Figure 2. Schematic representation of the structure of cilia. Schematic diagrams of cross sections of a typical primary cilium, a motile cilium and a basal body are shown in the right. In the middle is a schematic illustation of the primary cilium and the basal body. Also indicated are the transitional fibers and Y-link, all of which define the transition zone. In the primary cilium, Kinesin-2 motors drive anterograde transport of IFT particles and presumed cargoes (e.g. IFTdynein and tubulins) from the ciliary base to tip; at the tip, the IFT machinery is remodeled into retrograde and microtubules are constantly turning over; IFT-dynein motors recycle the IFT machinery and ciliary turnover products (e.g. tubulins) back to the ciliary base. Note that kinesinII and OSM-3 cooperate to drive anterograde transport in the middle segment (composed of outer doubles), whereas OSM-3 alone takes over the distal segment (composed of singlets) transport. The components of IFT particles, which are composed of motors, IFT-A and IFT-B subcomplexes, BBSome are listed in the left. Like centrioles, basal bodies are cylindrical structures of nine triplet microtubules. Each triplet consists of one complete 13-protofilament microtubule A fused to two incomplete microtubules B and C (11 protofilaments). The A and B tubules of the triplets extend to form outer doublets of the ciliary axoneme, whereas the C tubules terminate at the transition zone, a specialized region between basal body and axoneme (Lodish et al., 21 2000). In the transition zone, there are a series of structures visualized under electron microscope as Y-shaped linking the microtubule to the ciliary membrane (Gilula and Satir, 1972). These Y-links organize and terminate in “ciliary necklaces”, a specialized structural feature of the ciliary membranes (Gilula and Satir, 1972). Together these structures form a barrier to regulate ciliary entry and prevent random diffusion of both cytosol and membrane-bound macromolecules into the cilium. In many cilia the A tubules of the doublets extend as singlets in the distal segment (Silverman and Leroux, 2009) (Figure 2). The axonemal tubulins are subjected to posttranslational modifications including acetylation, detyrosination, polyglutamylation and polyglycylation (Janke and Bulinski, 2011) that appear to be important for the stability and function of cilia as suggested by loss-of-function studies on the modifying enzymes (Lee et al., 2012; Pathak et al., 2011; Pugacheva et al., 2007; Shida et al., 2010). The axoneme is about 0.25 µm in diameter, but the length varies from 1 to 10 µm (Lodish et al., 2000). The length is tightly controlled as discussed in detail in the section 4.4. The elongation of axoneme occurs by addition of tubulin subunits to the microtubule plus end at the ciliary tip. Interestingly, the microtubules are constantly turning over at the plus end and continuous incorporation of fresh subunits at the tip is needed to maintain the steady-state length of flagella in Chlamydomonas reinhardtii (Marshall and Rosenbaum, 2001). 4.2. IFT: the core machinery for cilium assembly and maintenance Because there is no protein synthesis machinery (ribosome) in cilia/flagella, ciliary precursors such as tubulins are synthesized in the cytoplasm and imported through a bidirectional transport system called “Intraflagellar transport system (IFT)”. IFT was first discovered microscopically as the bi-directional movement of granule-like particles along the flagella of Chlamydomonas reinhardtii (Kozminski et al., 1993). The studies on IFT in Chlamydomonas have greatly enhanced our understanding of cilia biology. IFT is an evolutionally conserved multiprotein complex (Table 1) transporting cargoes in (anterograde) and out (retrograde) of cilia/flagella between the outer doublet microtubules and the membrane. IFT cargoes include tubulins and other components for 22 ciliary structure as well as signaling molecules. IFT is required for assembly and maintenance of nearly all eukaryotic cilia and flagella (Plasmodium cilia and the sperm flagella of Drosophila are examples that assemble in the cytoplasm and do not require IFT (Pazour, 2004)). IFT dysfunction results in structural and functional defective cilia, causing system-wide disorders (Arts et al., 2011; Avasthi and Marshall, 2012; Bredrup et al., 2011; Davis et al., 2011; Gilissen et al., 2010; Mill et al., 2011; Perrault et al., 2012; Sung and Leroux, 2013) (Table 1). The anterograde IFT is powered by two types of plus-end directed motors in the kinesin-2 family: heterotrimeric kinesin-II and homodimeric Kif17/OSM-3. Kinesin-II consists of a heterodimer of two kinesin-2 motor subunits (Kif3a and Kif3b) and an accessory subunit, kinesin-associated protein (KAP). All three subunits are essential for the assembly and maintenance of cilia/flagella in most ciliated organisms (Kozminski et al., 1995; Marszalek et al., 1999; Mueller et al., 2005; Nonaka et al., 1998; Sarpal et al., 2003). In vertebrates, Kif3a can form a heterodimer with Kif3c instead of Kif3b. A redundancy between Kif3b and Kif3c has been suggested in a limited subset of cilia (Zhao et al., 2012). In C. elegans sensory cilia, kinesin-II cooperates with OSM-3 to drive the middle segment anterograde transport at an intermediary rate. At the distal end of middle segments, kinesin-II dissociates from IFT particles, which are then moved by OSM-3kinesin alone to the distal segment at a faster rate. A similar segmentation is also observed in photoreceptor cilia in zebrafish, where the OSM-3 homolog Kif17 builds distal singlets and targets visual pigment proteins to the outer segment (Insinna et al., 2008). In contrast to a general requirement of kinesin-II for ciliogenesis, Kif7 appears to be dispensable at least for renal cilia (Insinna et al., 2008). The tissue-specific requirements of the two types of kinesin motors perhaps reflect different distal structures in cilia of these tissues. The retrograde IFT is driven by cytoplasmic dynein-2, previously known as cytoplasmic dynein 1b (Ishikawa and Marshall, 2011). Depletion or mutation of components in this complex results in short and stumpy cilia/flagella accumulating IFT particles (Hou et al., 2004; Huangfu and Anderson, 2005; May et al., 2005; Ocbina et al., 2011; Pazour et al., 1999; Pazour et al., 1998; Porter et al., 1999; Signor et al., 1999). The accumulation is 23 consistent with a role of the motor in retrograde IFT. IFT complex can be grouped biochemically and genetically into three subcomplexes: IFT complex A (IFT-A), IFT complex B (IFT-B) and BBSome. Mutations in any subunit within one group phenocopy each other, indicating a cooperativity and requirement for a holocomplex for functional IFT (van Dam et al., 2013). The most plausible function of IFT complex is to link cargoes to the motor proteins. The predominance of predicted protein-protein interaction domains in IFT and BBS proteins suggests IFT complex as a large platform with multiple protein interaction sites for binding of motors and cargoes (Bhogaraju et al., 2013b). Biochemical analyses of purified IFT particles have identified six IFT protein subunits in IFT-A and 14 in IFT-B (Cole et al., 1998; Ishikawa and Marshall, 2011) (Table 1). The majority IFT proteins structurally resemble vesicular trafficking proteins in having βpropeller and/or tetratricopeptide repeat (TPR) domains (Sung and Leroux, 2013). A set of IFT proteins (IFT74, IFT81, IFT20, IFT54, IFT57) instead have coiled-coils mediating protein-protein interactions. IFT20 for example has been suggested to link kinesin-II to IFT-B (Baker et al., 2003). IFT74 and IFT81 have been shown to form a tubulin-binding module required for ciliogenesis (Bhogaraju et al., 2013a). An additional domain identified in IFT proteins is the small GTP-binding protein domain in Rab-like proteins IFT22 (Rabl2) and IFT27 (Rabl4) (Sung and Leroux, 2013). IFT22 is required for motility and fertility in mammalian sperms and seems to be restricted to motile cilia (Lo et al., 2012). IFT27 functions together with IFT25 not for ciliogenesis but rather to regulate transport of Hedgehog signaling molecules in mammalian cilia (Keady et al., 2012). In most cases, depletion or mutation of IFT-B subunits results in short or absent cilia (Houde et al., 2006; Huangfu and Anderson, 2005; Jonassen et al., 2008; Pazour et al., 2000), resembling defects associated with kinesin-2 motors. Therefor IFT-B is regarded to function in anterograde trafficking. In contrast, mutations in IFT-A subunits in some cases cause short bulbous cilia similar to disruption of cytoplasmic dynein 2 subunits (Huangfu and Anderson, 2005; Iomini et al., 2009; Liem et al., 2012; Ocbina et al., 2011; 24 Tran et al., 2008), suggesting a retrograde transport function. Additional IFT components have been identified independently of the conventional biochemical analyses and categorized into IFT complex A or B accessory proteins (Table 1) based on genetic and physical interactions (Ishikawa and Marshall, 2011). Such IFT accessory proteins may function as “adaptors” that enhance IFT selectivity (Mukhopadhyay et al., 2010; Ou et al., 2005). A functional IFT depends on proper assembly of IFT particles at the ciliary base, coordinated movement of IFT-A and IFT-B along the axoneme and turnaround from anterograde to retrograde transport at the ciliary tip. A suggested regulator for these events is a complex termed “BBSome” consisting of proteins encoded by genes mutated in Bardet-Biedl syndrome (BBS) (Ou et al., 2005; Ou et al., 2007; Wei et al., 2012). To date, 17 genes (BBS1-16 and BBS18) have been associated with Bardet-Biedl syndrome, a ciliopathy characterized by retinopathy, polydactyly, kidney cysts, obesity, hypogonadism and mental retardation (Sung and Leroux, 2013). The BBSome is a complex consisting of eight highly conserved BBS proteins BBS1, BBS2, BBS4, BBS5, BBS7, BBS8, BBS9 and BBIP10 (BBS18) (Jin et al., 2010). Additional BBS proteins (BBS6, BBS10 and BBS12) form a complex that assists BBSome assembly (Seo et al., 2010). The small GTPase BBS3 (Arl6) recruits the BBSome to form coat complexes on liposomes consistent with the resemblance of BBS subunits to coated vesicles on the structural level (Jin et al., 2010). The predominance of β-propeller or TPR domains in BBS proteins suggests that the BBSome might co-evolve with IFT proteins to increase the versatility and specificity of ciliary cargo transport (Sung and Leroux, 2013). BBS5 in turn contains two pleckstrin homology-like (PH-like) domains binding to phosphoinositides and is essential for ciliation (Nachury et al., 2007). Consistent with harboring these characteristic domains, the BBSome has been shown to promote cilia membrane biogenesis (Nachury et al., 2007) and function in vesicular transport of ciliary membrane proteins (Berbari et al., 2008a; Berbari et al., 2008b; Domire et al., 2011; Jin et al., 2010). BBIP10 is a subunit that colocalizes with the BBSome in cilia but not at centriolar satellites (Loktev et al., 2008). Interestingly, 25 BBIP10 is required for ciliogenesis, microtubule stability and tubulin acetylation (Loktev et al., 2008), properties not commonly seen in other BBSome subunits (Kulaga et al., 2004; Mykytyn et al., 2004; Nishimura et al., 2004; Zhang et al., 2013). This suggests that BBSome-bound BBIP10 may function to couple acetylation of axonemal microtubules and ciliary membrane growth (Loktev et al., 2008). Table 1. Components of the IFT machinery; adapted from (Ishikawa and Marshall, 2011). IFT system General name C. C. elegans H. sapiens Implicated component reinhardtii in cell cycle regulation Kinesin-2 Heterotrimeric FLA10 KLP-20 KIF3A* − kinesin-II FLA8 KLP-11 KIF3B* FLA3 KAP-1 KAP3/ KIFAP3 Homodimeric OSM-3/KLP-2 KIF17 Cytoplasmic Heavy chain DHC1B CHE-3 DYNC2H1* dynein-2/ Intermediate FAP133 DYCI-1 WDR34 cytoplasmic chain dynein 1b Light D1BLIC D2LIC/XBX-1 DYNC2LI1 Intermediate chain Light chain LC8/ DLC-1 DYNLL1 FLA14 IFT complex A IFT144 IFT144 DYF-2 WDR19* IFT140 IFT140 CHE-11 IFT140* IFT139 IFT139 ZK328.7 THM1/ TTC21B* IFT122/ IFT122/ DAF-10 IFT122/ IFT122A FAP80 WDR10* IFT121/ IFT121 IFTA-1 WDR35* IFT122B IFT43 IFT43 IFT43/ C14orf179* IFT complex A TLP1 TUB-1 TULP3 accessory IFT complex B IFT172 IFT172 OSM-1 IFT172 IFT88 IFT88 OSM-5 IFT88* Yes (Robert et al., 2007) IFT81 IFT81 IFT-81 IFT81 IFT80 IFT80 CHE-2 IFT80/ WDR56* IFT74/ IFT72 IFT74/ IFT-74 IFT74/ IFT72 IFT72 IFT70 IFT70/ DYF-1 TTC30A/ FAP259 TTC30B 26 IFT57 IFT54 IFT57 IFT54/ FAP116 CHE-13 DYF-11 - IFTA-2 IFT57 IFT54/ TRAF3IP1/ MIPT3 IFT52/ NGD5 IFT46/ C11orf60 IFT27/ RABL4 IFT25/ HSPB11 RABL5* IFT52 OSM-6 IFT46 IFT52/ BLD1 IFT46 IFT27 IFT27 - IFT25 - IFT20 IFT25/ FAP232 IFT22/ FAP9 IFT20 Y110A7A.20 IFT20 BBS1 BBS2 BBS4 FAP22 DYF13 BBS1 BBS4 DYF-3 DYF-13 BBS-1 BBS-2 F58A4.14 CLUAP1 TTC26 BBS1* BBS2* BBS4* BBS5 BBS7 BBS8 BBS9 BBIP10 BBS5 BBS7 BBS8 BBS9 - BBS-5 BBS-7 BBS-8 BBS-9 - BBS5* BBS7* BBS8* BBS9* BBIP10* Yes (Jonassen et al., 2008) − Yes (Hs) (Kim et al., 2004a) - IFT22 IFT complex B accessory BBSome DYF-6 Yes (Qin et al., 2007) - C. reinhardtii, Chlamydomonas reinhardtii; C. elegans, Caenorhabditi elegans; H. sapiens, Homo sapiens; BBIP10, BBSome-interacting protein of 10 kDa; CHE, chemotaxis abnormal; CLUAP1, clusterin-associated protein 1; DAF-10, abnormal dauer formation 10; DYF, abnormal dye-filling; FAP, flagellar-associated protein; FLA, flagellar-assembly impaired; HSPB11, heat shock protein βBIP IFTA, IFT-associated; KAP, kinesin-associated protein; KIF, kinesin family; KIFAP3, KIF-associated protein3; KLP, kinesin like protein; MIPT3, microtubule-interacting protein associated with TRAF3; OSM, osmotic avoidance abnormal; THM1, TTC-containing Hedgehog modulator; TLP1, tubby-like protein 1; TRAF3IP1, TRAF3-interacting protein 1; TTC, tetratricopeptide repeat; TUB1, tubby protein homologue 1; TULP3, tubby-like protein 3; WDR, WD repeat-containing; XBX-1, X-Box promoter element-regulated 1. Aliases of the same gene are separated by slash (/); Asterisk (*) indicates that mutations of the gene have been identified in human ciliopathies. 27 4.3. Bidirectional crosstalk between ciliogenesis and the cell cycle Primary cilia are generally assembled on quiescent cells or on proliferating cells in the G1 phase of the cell cycle and disassembled prior to mitosis. After mitosis, the daughter cell that inherits the old mother centriole is able to reassemble a primary cilium before its sister cell and this confers an asynchronous response to Hedgehog signaling shown in cultured mammalian cells and in vivo in mouse neuroepithelium (Anderson and Stearns, 2009; Piotrowska-Nitsche and Caspary, 2012). Notably, a recent study demonstrated that in the developing neocortex a remnant of the ciliary membrane remains attached to the mother centriole through mitosis and is preferentially inherited by the daughter cell retaining stem cell character (Paridaen et al., 2013). Interestingly, at late neurogenic stages, this remnant is often lost and so is the asynchronous cilium assembly in symmetric neurogenic divisions that give rise to two differentiating daughter cells, suggesting that inheritance of a ciliary membrane remnant underlies the asynchronous cilium formation and is involved in asymmetric regulation of cell fate between daughter cells (Paridaen et al., 2013). The reasons for this assembly-disassembly cycle are unclear, although it has been proposed that primary cilia may provide a means of sequestering the centriole for duplication and its subsequent function as the poles of the mitotic spindle. There appears to be a reciprocal regulation of cilia and the cell cycle. For example, cilia are lost in many cancer cells displaying hyperproliferation and cells deficient in ciliogenesis undergo abnormal cell division and cytogenesis. Some of the IFT components have been found to function in cell cycle control (indicated in Table 1). For example, reduced IFT27 causes incomplete or asymmetric cytokinesis in Chlamydomonas (Qin et al., 2007). Overexpression of IFT88 prevents the G1/S transition and depletion of IFT88 promotes cell cycle progression in mammalian cells (Robert et al., 2007). IFT88 is also implicated in regulating proper spindle orientation during cell division (Delaval et al., 2011). There are also other molecules besides IFT proteins implicated to regulate both ciliogenesis and the cell cycle. Aurora kinases known for their functions in mitosis also function in cilia/flagella disassembly. In mammalian cells, HEF1-dependent activation of 28 Aurora A promotes cilium disassembly during the early G1/S transition and a tubulin deacetylase HDAC6 has been demonstrated as one important effector for this ciliary function of Aurora A (Pugacheva et al., 2007). Chlamydomonas CALK (Chlamydomonas Aurora A like kinase) is required for flagellar shortening and its phosphorylation state is correlated with flagellar length (Luo et al., 2011; Pan et al., 2004). An inhibitory effect of cilia on cell cycle progression has been suggested by studies on two length regulators involved with dynein. Nde1 (nuclear distribution gene E homologue 1) is a centrosome protein associated with a cytoplasmic dynein light chain LC8/DYNLL1, depletion of which led to cilia elongation and a cilium-dependent delay in cell cycle re-entry (Kim et al., 2011b). Phosphorylation of Tctex-1 (T-complexAssociated-Testis-Expressed 1-Like 1), another cytoplasmic dynein light chain, accelerates cilium disassembly and S-phase entry (Li et al., 2011). In unciliated cells or cells lacking essential IFT components to construct cilia, the delay in G1/S transition by depletion of Ned1 or Tectx-1 is reverted, demonstrating that the observed inhibition of cell-cycle progression is dependent on the presence of cilia (Kim et al., 2011b; Li et al., 2011). Studies on NIMA (never in mitosis)-related kinases (NEKs) seem to link defects in ciliary length control also with premature chromosome separation and aneuploidy in vertebrates. These are defects associated with cell division often seen in cancer cells. For example, Nek8 mutations associated with the mouse model for juvenile cystic kidney (jck) disease lead to excessively long cilia and multinucleated cells (Liu et al., 2002b) as well as genome instability (Choi et al., 2013). The proteins involved in ciliary length control that also have been reported to regulate the cell cycle are indicated in Table 2. 4.4. Cilium formation and length control Formation of cilia Cilium formation in vertebrate cells follows a series of ordered steps. In early ciliogenesis, a Golgi-derived ciliary vesicle fuses with the mother centriole at the distal end, followed by outgrowth of ciliary microtubules, assembly of the transition zone, elongation of the 29 axoneme, and coordinated formation of ciliary membrane. The basal body appendages appear to play several important roles at the initial step (Wang et al., 2013), which might explain the asynchrony of cilium formation between sister cells. The distal appendage protein Cep164 is essential for the initial vesicular docking that initiates cilia membrane biogenesis (Schmidt et al., 2012). A few transition zone factors are pre-tethered to the centriole distal ends before ciliogenesis, for example Cep162, the targeting of which to the distal ends has been demonstrated to promote and restrict formation of transition zone specifically at the ciliary base (Wang et al., 2013). Although it is widely acknowledged that the formation of axoneme depends on IFT, IFT appears not required for the basal body docking and anchoring nor the transition zone formation (Reiter et al., 2012). The details in establishing transition zone and structures in this region (including ciliary necklace, transition fibers and Y-links) during ciliogenesis are not known. Two transition zone modules composed of proteins encoded by genes mutated in two related ciliopathies MKS (Meckel-Gruber syndrome) and NPHP (nephronophthisis) have been identified and are collectively required for cilium formation (Williams et al., 2011). In C. elegans, synergic depletion of one component from each module is required to inhibit ciliogenesis associated with impaired attachment of the basal body-transition zone to the membrane (Williams et al., 2011). Axonemal doublets elongate immediately adjacent to the site of transition zone assembly by addition of tubulins to the microtubule plus end. The elongation of the axoneme requires a coordinated expansion of the ciliary membrane involving vesicle fusion (Garcia-Gonzalo and Reiter, 2012). Consistent with the direct implementation of vesicle trafficking in ciliogenesis, several proteins involved in vesicle budding (AP-1 and Clathrin), targeting (Rab8, Rab11 and TRAPP), tethering (Exocyst) and fusion (SNAREs) have been reported to regulate ciliogenesis (Kaplan et al., 2010; Mazelova et al., 2009; Westlake et al., 2011; Yoshimura et al., 2007; Zuo et al., 2009). The regulation of synthesis of cilium-specific proteins when cells construct cilia is largely unknown. Two transcription factors, abnormal dauer formation 19 (DAF-19) (Phirke et al., 2011) in C. elegans and forkhead box J1 (FOXJ1) in vertebrate cells (Yu et al., 2008), 30 have been implicated in regulating expression of ciliary genes. The synthesized ciliary precursors must be transported into cilia from the cytoplasm and moved to the assembly site by IFT. The parameters of the IFT machinery such as size, speed and cargo selection are potential control points for ciliogenesis (Ishikawa and Marshall, 2011). The protein transport between the cytoplasm and cilium is regulated and involved in cilium-based signal transduction. Many of the signaling events take place at the ciliary membrane, whereas the mechanisms regulating the ciliary membrane targeting of signaling proteins are not well understood. The ciliary base acts as a selective filter, allowing passage of proteins with specific properties or associated with transporters (Garcia-Gonzalo and Reiter, 2012). Several ciliary targeting sequences have been identified within membrane proteins such as fibrocystin, somatostatin, and rhodopsin and are required for selectively target these proteins to the cilium (Berbari et al., 2008a; Follit et al., 2010; Tam et al., 2000). Three diffusion-barrier or gatekeeper complexes have been identified at transition zone (NPHP1, 4, and 8; B9/MKS-JBTS and septins) to maintain the specific protein composition of the ciliary membrane (Chih et al., 2012; Delous et al., 2009; Garcia-Gonzalo et al., 2011; Hu et al., 2010; Otto et al., 2010; Sang et al., 2011). The mature transition zone thus functions as a ciliary gate-keeper for membraneassociated proteins. Some proteins are prevented from entering the cilium by interaction with the actin cytoskeleton (Francis et al., 2011). The selected membrane proteins are guided along the ciliary axoneme by association with IFT-A and IFT-B (Pedersen and Rosenbaum, 2008). The association to the microtubule axoneme may regulate selective retention of membrane proteins in cilia (Francis et al., 2011). The removal of membrane proteins is likely mediated by endocytosis (Kaplan et al., 2012; Molla-Herman et al., 2010). Studying these mechanisms is critical for understanding how the distinct composition of ciliary membrane is maintained and how the dynamic ciliary transportation during signaling transduction is achieved. As described, cilium formation is a complex program of macromolecular synthesis and assembly that is subjected to extensive regulations at various stages. Despite the increasing number of proteins identified to regulate both processes, it still remains 31 unknown how the timing of cilium formation is coupled with the cell cycle. Ciliary length control Ciliary length appears to be tightly controlled as distinct cell types possess cilia of different ranges of length and the cilia of abnormal length are functionally defective (Avasthi and Marshall, 2012; Ishikawa and Marshall, 2011). Since cilia are dynamic organelles requiring a continuous supply for their maintenance, it is not surprising that ciliary length is modulated by the same control points for ciliary formation. Indeed, many molecules and pathways known to regulate cilia formation also modulate ciliary length, including Notch, molecules involved in actin dynamics and transcription factors controlling expression of ciliary genes. A full list of proteins reported to regulate ciliary length is in Table 2. Most of our knowledge about length control of cilia has come from studies on Chlamydomona flagella. Genetic screens have identified five genes mutated in Chlamydomonas resulting in long flagella (LF1-5), of which LF2, LF4 and LF5 encode protein kinases (Asleson and Lefebvre, 1998; Tam et al., 2013). LF2 is homologous to mammalian CCRK, localizes to cytoplasm and interacts with LF1 and LF3 that lack homologs in mammals (Tam et al., 2007). Notably, in contrast to the lf2 mutants representing partial loss of function of LF2p with long flagella, the complete loss of LF2 (the lf2-6 mutant) leads to short flagella (Tam et al., 2007), suggesting a dose dependent effect of LF2 on flagellar length. Depletion of the LF2 homolog in zebrafish instead results in curled cilia in kidney tubules (Ko et al., 2010), suggesting a context dependent effect of this conserved kinase in different ciliated organisms. LF4 is localized in both flagella and cytoplasm (Asleson and Lefebvre, 1998; Berman et al., 2003). LF4 shares the highest homolog with two related human kinases, intestinal cell kinase (ICK) and male germ cell-associated kinase (MAK). ICK and MAK belong to the family of ros cross-hybridizing kinase (RCKs) family within the CMGC (CDK/MAPK/GSK3/CLK) group of the human kinome. ICK and MAK contain a MAPK-like Thr-Asp-Tyr (TDY) motif in the activation loop essential for the kinase activity (Fu et al., 2006; Togawa et al., 2000; Wang and Kung, 2012). Interestingly 32 CCRK (the LF2 homolog) has been shown to activate ICK and MAK through phosphorylating threonine T157 of this motif (Fu et al., 2006; Wang and Kung, 2012). The biological relevance of this phosphorylation is not known and the genetic interactions between LF2 and LF4 remain elusive (Asleson and Lefebvre, 1998; Berman et al., 2003; Tam et al., 2007). Disruption of the LF4 homologs in Leishmania (LmxMPK9) and C. elegans (DYF-5) also results in long cilia (Bengs et al., 2005; Burghoorn et al., 2007). Mak-null mouse photoreceptor cells display longer cilia, and overexpression of MAK in NIH3T3 cells reduces ciliary length in a kinase-dependent manner (Omori et al., 2010). LF5 is homologous to the human cyclin-dependent kinase-like (CDKL) kinase, CDKL5 (Tam et al., 2013). Remarkably, LF5p localizes to the proximal end of flagella but translocates to the flagellar tip or along the flagella in lf1, lf2 and lf3 mutants (Tam et al., 2013), suggesting that the proximal localization of LF5p may be important for length control. The mechanisms by which these proteins regulate ciliary length are poorly known. The IFT rate appears to be increased in the lf4 mutant (Ludington et al., 2013). The C. elegans homolog Dyf-5 has been suggested to regulate the compartmentalization of the two types kinesin-2 motors, where Dyf-5 is required to restrict kinesin-II to the middle segments (Burghoorn et al., 2007). In the dyf-5 loss-of-function mutant, IFT proteins aggregate in the distal segments of the elongated cilia, suggesting a possible failure in switching from anterograde to retrograde transport (Burghoorn et al., 2007). It remains to be established whether these changes in IFT cause the elongation of cilia. The abnormal accumulation of certain IFT components in cilia has also been observed in C. elegans dyf-18 (CCRK homologue) mutant and in Mak-null mouse photoreceptor cells (Omori et al., 2010; Phirke et al., 2011). Together these observations suggest that CCRK and ICK/MAK may regulate ciliogenesis through IFT. The lack of mechanistic information about individual length regulator has prompted the development of theoretical models of length control as a combination of assembly and disassembly processes. Current models are again based on a few key observations from 33 Chlamydomonas flagella. The assembly of flagella occurs with decelerating kinetics (Ludington et al., 2013; Marshall et al., 2005; Marshall and Rosenbaum, 2001) and is driven by the injection of large anterograde IFT particle aggregates (IFT trains) during the early growth when much additional growth is required. As the length is approaching the steady-state level, it is fine-turned with injections of smaller IFT trains (Engel et al., 2009), although the total amount of IFT proteins is constant and independent of length (Marshall and Rosenbaum, 2001). The assembly slows as flagella elongate but remains active at the tip even after they reach the final length (Marshall, 2002; Marshall and Rosenbaum, 2001; Rosenbaum and Child, 1967). This steady-state assembly is balanced by the continuous removal of subunits from the tip (Ishikawa and Marshall, 2011; Marshall and Rosenbaum, 2001; Qin et al., 2004). When IFT is inhibited by mutating the anterograde motor, flagella immediately begin to shorten at a constant rate (Kozminski et al., 1995; Marshall et al., 2005; Marshall and Rosenbaum, 2001; Parker and Quarmby, 2003), implying that the disassembly rate at the tip is independent of ciliary length. These observations propose the “balance-point model”, where the steady-state length is determined by the balance of the assembly rate inverse of the length and the disassembly rate is constant. This model has been able to explain many experimental observations (Ishikawa and Marshall, 2011; Marshall et al., 2005), although a recent report has questioned whether the disassembly rate is always constant (Hilton et al., 2013). An alternative model for length control is that the activity of a reporter molecule directly related to the length is at the center of a feedback system integrating the regulations for assembly and disassembly. This model is stimulated by the observed changes in the phosphorylation status of Chlamydomonas CALK that has been mentioned in the section 4.3 (Cao et al., 2009; Cao et al., 2013; Luo et al., 2011; Pan et al., 2004). Although the phosphorylation and kinase activity of CALK is correlated with length, the substrates or the downstream effectors of CALK remain unknown (Cao et al., 2009; Cao et al., 2013; Luo et al., 2011; Pan et al., 2004). Further studies on this kinase might help to test the feedback model of length control. 34 Table 2. Proteins reported to regulate ciliary length. Class Protein Actin-related protein Adenylate cyclase Centrosomal protein Dynein GTPase activating protein Histone deacetylase Kinase ACTR3 (Hs) (Kim et al., 2010) ACIII (Mm, Hs) (Ou et al., 2009) Cep131 (Dr) (Wilkinson et al., 2009) Cep162 (Hs) (Wang et al., 2013) Cep290 (Mm) (Rachel et al., 2012) Cep70 (Dr) (Wilkinson et al., 2009) Nde1 (Mm, Hs) (Kim et al., 2011b) Tctex-1 (Mm, Hs) (Li et al., 2011) Bromi (Mm) (Ko et al., 2010) Tsc1 (Dr, Mm) (DiBella et al., 2009; Hartman et al., 2009; Yuan et al., 2012) Tsc2 (Mm) (Hartman et al., 2009) HDAC6 (Hs) (Pugacheva et al., 2007) Aurora-A (Hs) (Pugacheva et al., 2007) CALK (Cr) (Cao et al., 2013) GSK-3β (Cr, Dr) (Wilson and Lefebvre, 2004; Yuan et al., 2012) LF2p (Cr) (Tam et al., 2007); Dyf-18 (Ce) (Phirke et al., 2011); Cdk20 (Dr) (Ko et al., 2010); Cdk20 (Mm, Hs) (this study) Ciliary localization − Cilia Implicated in cell cycle regulation − − Ciliary base Yes (Hs) (Staples et al., 2012) − Centriole distal ends Transition zone Ciliary base Mother centriole Transition zone − Basal body − Cilia Basal body Basal body Flagella No LF4p (Cr) (Berman et al., 2003); MPK9 (Lm) (Bengs et al., 2005); Dyf-5 (Ce) (Burghoorn et al., 2007); ICK (Mm, Hs) (this study) LF5p (Cr) (Tam et al., 2013) Flagella/Cilia MAK (Mm, Hs) (Omori et al., Cilia distal end Flagella − Yes (Hs) (Shi et al., 2011) Yes (Mm, Hs) (Kim et al., 2011b) Yes (Mm, Hs) (Li et al., 2011) − Yes (Mm) (WatayaKaneda et al., 2001) Yes (Mm) (WatayaKaneda et al., 2001) − Yes (Hs) (Fu et al., 2007) − Yes (Hs) (Diehl et al., 1998) Yes (Hs) (An et al., 2010; Feng et al., 2011; Liu et al., 2004; Ng et al., 2007; Wohlbold et al., 2006) and this study Yes (Hs) (Fu et al., 2009) and this study Yes (Hs) (Valli et al., 2012) Yes (Hs) (Wang and 35 2010) and this study Kinesin MT associate protein Phosphatase Phospholipase Receptor Scaffolding protein Small GTPase Kung, 2012) − Nek1 (Mm) (Shalom et al., 2008) Nek4 (Hs) (Coene et al., 2011) Nek8/NPHP9 (Mm) (Manning et al., 2013) S6k1 (Dr) (Yuan et al., 2012) Basal body KLP-6 (Ce) (Morsci and Barr, 2011) DCDC2 (Rn)(Massinen et al., 2011) RP1 (Mm) (Omori et al., 2010) Cdc14b (Dr) (Clement et al., 2011) PTPN23 (Hs) (Kim et al., 2010) PLA2G3 (Hs) (Kim et al., 2010) Fgfr1 (Dr, Xl) (Neugebauer et al., 2009) Cilia − Yes (Hs)(Choi et al., 2013) Yes (Hs) (Fingar et al., 2004) − Cilia − Cilia distal end No Basal body − Yes (Hs) (Mocciaro and Schiebel, 2010) − Pericentrosome − − HEF1 (Hs) (Pugacheva et al., 2007) Arl-13 (Ce) (Li et al., 2010) Arl-3 (Ce) (Li et al., 2010) Basal body Cilia Ciliary base Yes (Hs) (Dombrowski et al., 2013) Yes (Hs) (Law et al., 1998) − Yes (Hs)(Zhou et al., 2006) − − − − No − Transition zone − Transition zone − Transition zone − Transition zone − Transition zone − − − Rab8 (Hs) (Nachury et al., 2007) Rab11 (Hs) (Knodler et al., 2010) Transcription factor DAF-19 (Ce) (Phirke et al., 2011); RFX3 (Mm) (Bonnafe et al., 2004) Foxj1 (Dr, Mm, Hs) (Cruz et al., 2010; Yu et al., 2008) Transition zone Cc2d2a (Mm) (Garcia-Gonzalo protein et al., 2011) NPHP8 (Hs) (Patzke et al., 2010) Tctn1 (Mm) (Garcia-Gonzalo et al., 2011) Tctn2 (Mm) (Garcia-Gonzalo et al., 2011) Tmem67 (Mm) (Garcia-Gonzalo et al., 2011) Transmemebrane Notch ligand DeltaD (Dr) protein (Lopes et al., 2010) MT, microtubule; ATCR3, actin-related protein 3; ACIII, Ciliary base Cilia − Doublet segment Cilia adenylyl cyclase III; Cep131, centrosomal 36 protein 131 kDa; Cep70, centrosomal protein 70 kDa; Nde1, nudE neurodevelopment protein 1; Tctex-1 (or DYNLT), dynein light chain Tctex-type 1; Bromi, broad-minded; Tsc, tuberous sclerosis complex; HDAC6, histone deacetylase 6; CALK, Chlamydomonas aurora-like protein kinase; GSK3β, glycogen synthase kinase 3; LF, long flagella; Dyf, dye filling-defective; MPK9, putative mitogen-activated protein kinase 9; Nek, NIMA (never in mitosis gene a)-related kinase; S6k1, ribosomal protein S6 kinase beta-1; KLP-6, kinesin-like protein 6; DCDC2, doublecortin domain containing 2; RP1, retinitis pigmentosa 1; Cdc14b, cell division cycle 14B; PTPN23, protein tyrosine phosphatase non-receptor type 23; PLA2G3, phospholipase A2 group III; Fgfr1, fibroblast growth factor receptor 1; HEF1, human enhancer of filamentation protein 1; Arl, adenosine diphosphate ribosylation factor–like protein; RFX3, regulatory factor X 3; Cc2d2a, coiled-coil and C2 domain containing 2A; NPHP8, ephrocystin 8; Tctn, tectonic; Tmem67, transmembrane protein 67; DeltaD, delta-like protein D. Cr, Chlamydomonas reinhardtii; Ce, Caenorhabditis elegans; Dm, Drosophila melanogaster; Dr, Danio rerio; Hs, Homo sapiens; Lm, Leishmania mexicana; Mm, Mus musculus; Rn, Rattus norvegicus; Xl, Xenopus laevis. 4.5. The cilia-mediated signaling and links to cancer A number of critical cell signaling molecules localize in primary cilia and require an intact cilium for signaling transduction. These include components of the Hedgehog (Hh), the canonical Wingless-Int (Wnt) (Lancaster et al., 2011) and platelet derived growth factor receptor α (PDGFRα) pathways (Schneider et al., 2005). The role of cilia in mediating Hh signaling has been characterized most extensively. Hh signaling is important for cell proliferation and embryonic patterning. The mouse models carrying mutations in the IFT machinery display characteristic Hh signaling defects in neural tube patterning and limb development (Ashe et al., 2012; Garcia-Garcia et al., 2005; Huangfu and Anderson, 2005; Ko and Park, 2013; Liem et al., 2012; Marszalek et al., 1999). Genetic and biochemical analyses have revealed that these phenotypes are associated with disruption of Hh signaling, demonstrating that an intact cilia is required for the Hh signaling activation in mammals. The Hh receptor Patched 1 (Ptch) localizes in cilia in the absence of ligand (Rohatgi et al., 2007). Upon ligand binding, the G protein coupled receptor Smoothened (Smo) subsequently translocates to cilia and stimulates the Gli family of transcription factors (Tukachinsky et al., 2010). There are three mammalian paralogs of the Gli family members: Gli1 is a transcriptional activator unregulated after initial pathway activation; 37 Gli2 and Gli3 can act as activators or repressors depending on proteolytic processing (Pan et al., 2006; Wen et al., 2010). In the absence of ligand, Gli2 and Gli3 interact with negative regulators Sufu and Kif7 as a scaffold for the kinases promoting a proteolytic cleavage of Gli2 and Gli3 into the truncated repressors (Chen et al., 2009; Tukachinsky et al., 2010). Upon ligand induction, Kif7, Sufu, Gli2 and Gli3 translocate to the ciliary tip and the interaction of Sufu with Gli is lost, allowing stabilization of full-length Gli (Tukachinsky et al., 2010). Although Sufu regulates Gli independent on cilia (Chen et al., 2009), the proteolytic processing of Gli is inhibited in cells with deficient cilia (Huangfu and Anderson, 2005; Humke et al., 2010). The cilium-dependent formation of Gli repressors suggests a negative effect of cilia on Hh signaling in the absence of ligand. The activation mechanism of full-length Gli by Smo in cilia is not known but translocation of Smo to cilia is required for this activation (Tukachinsky et al., 2010; Wen et al., 2010; Wu et al., 2012). The level of Gli2 and Gli3 enriched at the tips correlates with increased responsiveness (Chen et al., 2009; Tukachinsky et al., 2010; Wen et al., 2010). The activated Gli is transported out cilia and enters the nucleus to drive expression of Gli1 and other Hh target genes. Many known Hh target genes are involved in cell growth and proliferation (Varjosalo and Taipale, 2008), including key regulators of G1/S transition- cyclin D and cyclin E (Duman-Scheel et al., 2002). Because the involvement of cilium in multiple mitogenic signaling pathways and its implication in cell cycle progression, it has been proposed that defective ciliogenesis could represent an important step in tumorigenesis. Two elegant genetic studies in mice have revealed an unexpected dual role of cilia in the development of the basal cell carcinoma (BCC) and medulloblastoma driven by mutations in the Hh signaling components (Han et al., 2009; Wong et al., 2009). Expression of a constitutively active Smo in the postnatal mouse brain causes medulloblastoma and ablation of cilia strongly inhibits tumor formation associated with decreased proliferation (Han et al., 2009), consistent with earlier genetic studies demonstrating that cilia are required for Hh signaling activation at a step downstream of Smo (Huangfu and Anderson, 2005). Constitutively active Gli2 can also cause medulloblastoma in mice if cilia are removed, demonstrating that cilia depletion is required for medulloblastoma formation driven by 38 the Gli2 mutation (Han et al., 2009). Similar results were obtained in the mouse model for BBC driven by the same mutations of Smo and Gli2 (Wong et al., 2009). These results suggest that primary cilia can be either permissive or suppressive for tumor formation, depending on the underlying oncogenic events in the upstream or downstream of Hh signaling. The opposing effects of cilia on the development of these tumors are likely reflecting the dual effects of cilia on Hh signaling. So far, a complete examination of cilia frequency in cancer cells compared to the corresponding normal tissues has not been performed. The analysis of the presence of cilia and mutations in the Hh components in human medulloblastomas and mouse BBCs is consistent with the prediction that tumors relying on upstream activation of Hh signaling retain cilia (Han et al., 2009; Wong et al., 2009). A decrease in cilia frequency has been documented in glioblastoma cell lines (Moser et al., 2009), breast cancer (Yuan et al., 2010), melanoma (Kim et al., 2011a), pancreatic cancer (Seeley et al., 2009) and ovarian cancer (Egeberg et al., 2012). The contributions of cilia depletion to these malignancies and the involvement of Hh signaling or other ciliary signaling remain to be established. Recent reports demonstrate that depletion or inhibition of HEF1, Aurora A or HDAC6 can restore cilia in renal cancer cells (Xu et al., 2010) and ovarian adenocarcinoma cells (Egeberg et al., 2012). However, it is not known whether the negative effects of HEF1, Aurora A and HDAC6 on cilia stability (Pugacheva et al., 2007) mentioned in the section 4.3 contribute to their oncogenic properties. Mutations in VHL (von Hippel-Lindau) tumor suppressor cause renal cysts that lack cilia and precede renal carcinogenesis (Lutz and Burk, 2006). Although restoration of the wild-type VHL is able to reverse the arrest of ciliogenesis (Lutz and Burk, 2006), it is not clear whether preservation of cilia is part of the tumor-suppressive function of VHL. CCRK is specifically overexpressed in glioblastoma cells and has been implicated in cell cycle regulation, but the reported molecular mechanisms appear controversial (An et al., 2010; Feng et al., 2011; Liu et al., 2004; Ng et al., 2007; Wohlbold et al., 2006). The observations that the homologs of CCRK are involved in regulating cilia/flagellar 39 morphology (Ko et al., 2010; Phirke et al., 2011; Tam et al., 2007) have prompted us to explore whether CCRK regulates ciliogenesis in mammalian cells and whether this function contributes to its oncogenic properties. 40 AIMS OF THE STUDY This study was to investigate the cellular functions of mammalian Cdk7 and CCRK kinases. The specific aims were: 1. To investigate the possible role of the Cdk7 kinase subunit of TFIIH basal transcription factor complex in the general production of mRNA and rRNA; 2. To investigate the possible role of Cdk7 in regulating the adipogenic transcriptional program mediated by the master adipogenic transcription factor PPARγ; 3. To investigate the potential role of CCRK in regulation ciliogenesis and the cell cycle. 41 MATERIALS AND METHODS The materials and methods used in this study are listed below and described in detail in the original publications referred to by Roman numerals. 1. Materials Cell lines 293-shhN Description HEK 293 cell line stably transfected with Shh-N expression and neomycin resistance constructs human embryonic kidney cells transformed with the SV40 large T antigen a pre-adipogenic substrain of mouse 3T3 derived from human 293 cells for the Flp-In T-Rex system (Invitrogen) to generate tretracycline-induciable cells line expressing gene of interest mouse embryonic fibroblasts immortalized with a C-terminal fragment of p53 mouse embryonic fibroblasts human glioblastoma cells human osteosarcoma cells Source Taipale lab (Chen et al., 2002) Used in IV Life Technologies IV ATCC III Life Technologies II Mäkelä lab I-III ATCC ATCC ATCC IV IV III Description Source or reference Used in Cdk2 fused to GST (Hermand et al., 1998) III Gateway entry clone Hs_ICK(CDS)/pEN TR human ICK, Ultimate ORF Clone IOH38087 Life Technologies IV Mm_CCRK (CDS)_si8_resist/pD ONR221 gateway entry clone of mouse Ccrk cDNA resistant to siCCRK8 (J-053874-08) generated by the author IV Mm_CCRK_(CDS)/ pDONR221 mouse Ccrk cDNA gateway entry clone, ID 38901 MGC Gene collection IV Taipale lab (Taipale et al., 2000) IV 293FT 3T3-L1 Flp-In 293 T-Rex Immortalized MEFs NIH3T3 U251MG U2OS Plasmids E. coli expression GST-Cdk2 Mammalian expression Gli-luc 8 Gli-binding sites driven expression of firefly luciferase reporter 42 Hs_ICK(CDS)_T15 7A/pDEST27 Hs_ICK(CDS)_T15 7E/pDEST27 Hs_ICK(CDS)/pDE ST27 Mm_CCRK (CDS)_si8_resist/pc DNA6.2_N-EmGFP Mm_CCRK_(CDS)/ pcDNA6.2_si8_resis t_K33M/ pcDNA6.2_NEmGFP pCMV-SPORT6PPARγ2 PPRE3-TK-Luc pRL-TK pSV-SPORTPPARγS112A shControl/pENTRH1 shRNA-K7/pENTRH1 Flp-In T-Rex system pOG44 Mm_Cdk7/pTO_HA _StrepIII_GW_FRT Mm_Cdk7_K41M/p TO_HA_StrepIII_G W_FRT CMV promoter-driven expression of human ICK T157A mutant, Nterminal GST-tag CMV promoter-driven expression of human ICK T157E mutant, Nterminal GST-tag CMV promoter-driven expression of human ICK, N-terminal GSTtag CMV promoter-driven expression of mouse Ccrk WT protein with silent mutations resistant to siCCRK-8 (J-053874-08), Nterminal GFP-tag CMV promoter-driven expression of mouse Ccrk K33M with silent mutations resistant to siCCRK-8 (J-053874-08), N-terminal GFPtag CMV promoter-driven expression of wild type PPARγ2 generated by the author IV generated by the author IV generated by the author IV generated by the author IV generated by the author IV MGC Gene collection III 3 PPRE upstream of the thymidine kinase promoter fused to luciferase the thymidine kinase promoterdriven renilla luciferase reporter plasmid CMV promoter-driven expression of S112 non-phosphorylated PPARγ2 mutant non-targeting shRNA, ATCCAAAAGCTGTCTTCGTT CTGCAG shRNA targeting human Cdk7, CCAATAGAGCTTATACAC Dr. Chatterjee III Promega IV Dr. Spiegelman III BCH Genome Biology Unit III BCH Genome Biology Unit III Flp recombinase expression plasmid Life Technologies (O'Gorman et al., 1991) II Expression vector containing mouse Cdk7 WT, N-terminal HA and StrepIII-tag Expression vector containing mouse Cdk7 K41M, N-terminal HA and StrepIII-tag generated by the author II generated by the author II TRC-Hs1.0 (Human) IV pseudotyped lentivirus shRNA system Hs_shIFT20_5/pLK shRNA targeting human IFT20, O.1 TRCN0000043375 43 Hs_shIFT20_6/pLK O.1 shRNA targeting human IFT20, TRCN0000043376 TRC-Hs1.0 (Human) IV Hs_shKIF3A_2/pLK O.1 shRNA targeting human KIF3A, TRCN0000116812 TRC-Hs1.0 (Human) IV Hs_shKIF3A_4/pLK O.1 shRNA targeting human KIF3A, TRCN0000116814 TRC-Hs1.0 (Human) IV Hs_shKIF3A_5/pLK O.1 shRNA targeting human KIF3A, TRCN0000116815 TRC-Hs1.0 (Human) IV Mm_shIft172/pLKO .1 shRNA targeting mouse Ift172, TRCN0000079815 TRC-Mm1.0 (Mouse) IV Mm_shKif3a_3/pLK O.1 shRNA targeting mouse Kif3a, TRCN0000090403 TRC-Mm1.0 (Mouse) IV Mm_shKif3a_3/pLK O.1 shRNA targeting mouse Kif3a, TRCN0000090405 TRC-Mm1.0 (Mouse) IV shNT/pLKO.1 Hs_shCCRK/pDSLhpUGIH non-targeting shRNA, SHC002 shRNA targeting human CCRK, generated from NA2.F gateway entry clone non-targeting shRNA, generated from gateway entry clone packaging plasmid used for pseudotyped lentivirus production. CMV-driven expression of Gag-Pol VSV-G (vesicular stomatits virus G-protein) expressing envelope plasmidused for pseudotyped lentivirus production Sigma generated by the author IV IV generated by the author IV (Moffat et al., 2006) IV (Moffat et al., 2006) IV shNT/pDSLhpUGIH pCMV-dR8.91 VSV-G The siRNAs used in this study are from Dharmacon, Thermo Scientific and listed below. siRNA ID J-053874-06 J-053874-07 J-053874-08 J-049932-05 J-049932-07 J-004686-06 J-004686-09 J-004811-09 J-004813-09 J-040601-05 J-040601-07 J-003241-12 Targeting gene mouse Ccrk mouse Ccrk mouse Ccrk mouse Ick mouse Ick human CCRK human CCRK human ICK human MAK mouse Cdk7 mouse Cdk7 human CDK7 Target sequence AAACGACAUUGAACAACUG GGAGAGCUGUUGAAUGGGU UGUGCUGGCUUUCGAAUUU GUGCAUACCCAAUAACUUA GGACUUUCUUCCAUCAAAU UCACUGAGCUGCCGGACUA AGGCACAGGUCAAGAGCUA GCACUUCGAUAUCCUUACU CAAGUGAGGUCGAUGAAAU GGACAUAAGUCUAACAUUA UGUGUAGUCUUCCCGAUUA GAUGACUCUUCAAGGAUUA Used in IV IV IV IV IV IV IV IV IV II II II 44 D-001810-01 D-001810-02 D-001810-03 Antigens acetylated α-tubulin BrdU Cdc2-T161-P Cdk2 Cdk2-T160-P Cdk7 Cdk7 GFP Gli3 GST H3K4trime IFT88 Mat1 PPARγ PPARγ-S112-P RNA Pol II RNA Pol II RNA Pol II-S5-P RNA Pol II-S5-P streptavidin-HRP γ-tubulin ON-TARGETplus Non-Targeting #1 ON-TARGETplus Non-Targeting #2 ON-TARGETplus Non-Targeting #3 UGGUUUACAUGUCGACUAA II-IV UGGUUUACAUGUUGUGUGA II, IV UGGUUUACAUGUUUUCUGA II, IV Description T7451, mouse monoclonal Ab B2531, mouse monoclonal Ab #9114, rabbit polyclonal Ab sc-163, rabbit polyclonal Ab #2561, rabbit polyclonal Ab C-4, mouse monoclonal Ab C-19, rabbit polyclonal Ab TP401, rabbit polyclonal Ab AF3690, goat polyclonal Ab ab9085, rabbit polyclonal Ab ab8580, rabbit polyclonal Ab 13967-1-AP, rabbit polyclonal Ab FL-309, rabbit polyclonal Ab ab41928, mouse monoclonal Ab IF5, mouse monoclonal Ab ab5408, mouse monoclonal Ab N20, rabbit polyclonal Ab H14, mouse monoclonal Ab ab5131, rabbit polyclonal Ab P0397, peroxidase-conjugated T5192, rabbit polyclonal Ab Source Sigma Sigma Cell Signaling Santa Cruz Cell Signaling Santa Cruz Santa Cruz Torrey Pines Biolabs R&D systems Abcam Abcam Proteintech Santa Cruz Abcam Euromedex Abcam Santa Cruz Nordic Biosite Abcam Dako Sigma Used in IV II, III III III III I-III II IV IV IV II IV I-III III III II I-II I-II I-II II IV 2. Methods Method 4sU labeling Agarose gel electrophoresis Biotinylation and purification of 4sU-labeled RNA Cell culture ChIP Crystal violet staining Detection of 4sU-labeled RNA by RT-qPCR Detection of 4sU-labeled RNA by steptavidin-HRP blot EdU labeling and staining EU labeling and staining Flow cytometry analysis Used in II I-IV II I-IV I-II IV II II IV I IV 45 Flp-In 293 T-Rex cell line generation FUrd labeling and staining Gateway cloning Immunofluorescence analysis (IF) Luciferase reporter assay Microscopy Mutagenesis using QuickChange Lightning Site-Directed and Multi Site-Directed Mutagenesis Kit (Agilent) plasmid purification using GenElute Miniprep kit (Sigma) plasmid purification using NucleoBond midiprep kit (Macherey-Nagel) pseudotyped lentivirus infection pseudotyped lentivirus production Reverse transcription and RT-qPCR RNA extraction using RNeasy Plus Mini Kit (Qiagen) RNA extraction using TRI Reagent (MRC, TR118) siRNA transfection Transfection of cells Western blot analysis II I-II II, IV I-IV III, IV I-IV II, IV I-IV I-IV IV IV I-IV I-IV I-IV I-IV I-IV I-III The main methods used by the author are described below in detail. Gateway cloning and mutagenesis (II, IV) A mouse Cdk7 cDNA clone (EST cDNA IMAGE ID 479857) was used to amplify the Cdk7 open reading frame using primers with flanked attB sites and the resulting PCR product was inserted to pDONR221 (Invitrogen) to generate Mm_Cdk7_WT/pDONR221. A mutation was introduced to generate a kinase-deficient Cdk7 encoding plasmid Mm_Cdk7_K41M/pDONR221 using Mm_Cdk7_K41M_F and R primers with QuickChange Site-Directed Mutagenesis Kit (Agilent). Sequence-verified entry clones were transferred to Gateway destination vector pTO_HA_StrepIII_GW_FRT (Glatter et al., 2009) according to manufacturer ́s protocol (Invitrogen). A mouse Ccrk cDNA clone (MGC ID 38901; BC031907) was used to amplify the Ccrk open reading frame using primers with flanked attB sites and the resulting PCR product was inserted to pDONR221 (Invitrogen) to generate Mm_CCRK_(CDS)/pDONR221. Seven silent mutations within the target region of siRNA J-053874-08 were subsequently introduced into this plasmid using the primer Mm_Ccrk_si08_resis_mut and QuickChange Lightning Multi Site-Directed Mutagenesis Kit (Agilent) to generate Mm_CCRK_(CDS)_si8_resist/pDONR221. A further mutation was introduced to generate a kinase-deficient CCRK encoding plasmid Mm_CCRK_(CDS)_K33M 46 si8_resist/pDONR221 using Mm_Ccrk_K33M_F and R primers with QuickChange SiteDirected Mutagenesis Kit (Agilent). Sequence-verified entry clones were transferred to pcDNA6.2/N-EmGFP (Invitrogen) according to manufacturer’s protocol. Similarly, Hs_ICK(CDS)/pDEST27 was generated by transferring the human ICK Ultimate ORF Clone (IOH38087; Invitrogen) to pDEST27 (Invitrogen). Generation of Hs_ICK(CDS)_T157A/pDEST27 and Hs_ICK(CDS)_T157E/pDEST27 was as described above with the following primers: Hs_ICK_T157A_F2, Hs_ICK_T157A_R2, Hs_ICK_T157E_F and Hs_ICK_T157E_R and verified by sequencing. Primer sequences are in II and IV. siRNA transfection (II, IV) 1.5-3 x 105 NIH3T3 and U251MG cells were seeded per well of a 6-well plate (IV) or 810 x 105 immortalized MEFs (II) per 15 cm plate with DMEM supplemented with 10% Fetal Bovine Serum (FBS, Gibco) and 1% L-glutamine one day before transfection. 5 µl of 20 µM siRNAs and 5 µl Lipofectamine 2000 or RNAiMAX (Invitrogen) were mixed with 250 µl DMEM separately. After 5 min, the siRNA and RNAiMAX dilutions were mixed by vortexing and incubated for 20 min. The cells were changed into 1.5 ml DMEM supplemented with 10% FBS, 1% L-glutamine per well of a 6-well plate or 4 ml per 15 cm plate before adding the transfection mixture. 4 ml DMEM supplemented with 10% FBS, 1% L-glutamine was added to each 15 cm plate 7 h later and replaced by 20 ml DMEM supplemented with 10% FBS, 1% L-glutamine, 100 U/ml penicillin G, 100 µg/ml streptomycin at 24 h. The siRNA transfection was performed again with a 24 h interval when indicated. In experiments with double and triple siRNA transfections, control (siNT) transfections were performed with all concentrations. Immunofluorescence Staining (I-IV) Cells grown on glass coverslips were fixed at indicated time points with 4% paraformaldehyde for 15 min. To visualize γ-tubulin, the coverslips were incubated in PHEM buffer (1% Triton, 45 mM HEPES pH 7.2, 45 mM PIPES, 10 mM EGTA, 20 mM MgCl2) for 45 sec to 1 min before fixation. After a PBS wash, cells were permeabilized with PBS-0.25% Triton for 20 min. The coverslips were blocked with 10% goat or horse serum in PBS-0.25% Triton for 30 min and incubated with indicated primary antibodies 47 for 1 h. The coverslips were then washed three times with PBS followed by incubation with secondary antibodies for 30 min in the dark. Secondary antibodies (Life Technologies) used were donkey anti-mouse Alexa 488 (1:500), donkey anti-goat Alexa 594 (1:500), goat anti-rabbit Alexa 594 (1:1000), goat anti-mouse Alexa 594 (1:1000), goat anti-mouse Alexa 647 (1:500), goat anti-rabbit Alexa 647 (1:500), goat anti-rabbit Alexa 488 (1:500), goat anti-mouse Alexa 488 (1:500), and rabbit anti-mouse Alexa 488 (1:500). The washing step was repeated and Hoechst staining was done with 0.5 mg/ml of Hoechst H33342 (Sigma) for 5 min prior to the final wash, after which the stained coverslips were mounted onto glass slides and imaged using Zeiss Axioplan 2 microscope. In III, the z-stacks images were taken using a 63X oil objective and Axiovision 4.8 software to produce maximum intensity projection (MIP) images for ciliation and length measurement. Chromatin immunoprecipitation (I-II) 1-2 x 106 MEFs in one 15 cm plate were cross-linked with 9.5 ml of 1% formaldehyde diluted in PBS for 15 min at room temperature. The cross-linking was terminated by adding 0.5 ml of 2.5 M glycine in PBS and kept for 5 min on shaker. Fixed cells were lysed with 4 ml of RIPA buffer including inhibitors (150 mM NaCl, 1% NP40, 1% sodium deoxycholate, 1% SDS, 50 mM Tris pH 8.0, 3 mM EDTA, 1 mM DTT, 0.25 µg/ml Aprotinin, 0.5 mM PMSF, 10 mM β-Glycerophosphate, 10 mM NaF, 1 µg/ml leupeptin) after 2 washes with 7 ml cold PBS. The cell lysates were collected using cell scrapper and sonicated using Branson Sonifier cell disruptor B15 (output control level 5, 30% duty cycle, 15 sec x 10, 1 min rest on ice between). The sonicated lysates were precleared with 40 µl of 50 % slurry of protein-A sepharose (PAS, Sigma, P3391) for 2 h with rotation at 4°C. PAS was saturated with 0.3 µg/ml Herring Sperm DNA for 2 h with rotation at room temperature followed by 3 washes with RIPA buffer and made into 50% slurry. For immunoprecipitation, 1 mg of the pre-cleared lysates were incubated with 50% slurry of saturated PAS and 2-4 µg of indicated antibodies or mouse IgG (Sigma, I5381) with rotation overnight at 4°C. The beads were washed twice with 1 ml RIPA buffer, four times with 1 ml Szak’s IP wash buffer (100 mM Tris pH 8.5, 500 mM LiCl, 1% NP40, 1% sodium deoxycholate) and twice with 1 ml RIPA buffer including a 5 min rotation at 4°C between each wash. After another 2 x wash with 1 ml cold TE buffer (10 mM Tris pH 7.5, 48 1 mM EDTA), 200 µl 1.5 x Talianidis elution buffer (70 mM Tris pH 8.0, 1 mM EDTA, 1% SDS) were added to the remaining 100 µl TE buffer and beads mix and the immunocomplexes were eluted for 10 min at 65°C. The cross-linking was reversed by adjusting to 200 mM NaCl and incubating 5 h at 65°C. DNA was purified using QIAquick PCR purification kit (Qiagen) and was used as template in real-time PCR reactions. RNA extraction using TRI Reagent (I-II) 5 ml Trizol (MRC, TR118) was added to cells per 15 cm plate after 2 washes with 7 ml PBS and collected into a 15 ml falcon tube using cell scrapper. After 5 min, 1 ml of chloroform was added and vigorously shaken with hand for 15 sec. After 3 min, the samples were centrifuged at 5000 rpm at 4°C for 15 min. The upper aqueous phase was transferred to a fresh tube with 2.5 ml isopropanol and mixed well. After 10 min of incubation, RNA was precipitated by centrifugation at 5000 rpm at 4°C for 10 min. The pellet was washed with 5 ml 75% ethanol and air dried in a fume hood. The pellet was then dissolved in 100-300 µl RNAse-free water (Thermo), incubated at 60°C for 10 min and stored at − 20°C till use. Reverse transcription and RT-qPCR (I-IV) If no special indications, total RNA was isolated using RNeasy Plus Mini Kit (Qiagen) according to manufacturer’s protocol. 400 ng of total RNA was used as template in 20 µl cDNA reactions with TaqMan MULV reverse transcriptase and random hexamer or OligodT as indicated (Applied Biosystems). 1 µl of the cDNA reaction was used as template in a 20 µl RT-qPCR reaction with SYBR-green RT-PCR reagent (Applied Biosystems) and 200 nM final concentration of forward and reverse primers (Sigma, sequences in I-IV). The comparative threshold cycle (CT) method was used to analyze relative expression curves on a StepOnePlus Real-Time PCR System (Applied Biosystems). 4sU labeling and detection by streptavidin-HRP blot (II) For metabolic labeling of RNA in MEFs, 4-thiouridine (4sU, Sigma, T4509) was applied to cells at a 200 µM final concentration in pre-warmed media for 30 min. Total RNA was 49 isolated using TRI reagents, of which 15-50 µg was used for the biotinylation reaction as previously described (Dolken et al., 2008). Biotinylation of 4sU-labeled RNA was performed at a final concentration of 100 ng/µl RNA in 10 mM Tris pH 7.5, 1 mM EDTA, and 0.2 mg/ml Biotin-HPDP (Pierce) for 1.5 h at room temperature in dark. The unbound Biotin-HPDP was removed by chloroform/isoamylalcohol (24:1) extraction using Phase-lock-gel (Heavy) tubes (5Prime) and RNA was precipitated by isopropanol followed by a wash with 75% ethanol. The pellet was resuspended in 40 µl RNAse-free water (Thermo) and denatured at 65°C for 10 min followed by rapid cooling on ice for 5 min. RNA samples were mixed with 10 µl of 5 x RNA loading buffer (16 µl saturated aqueous bromophenol blue solution, 80 µl 500 mM EDTA pH 8.0, 720 µl 37% formaldehyde, 2 ml 100% glycerol, 3084 µl formamide, 4 ml 10 x MOPS, RNase-free water to 10 ml) and 0.5 µl of 5 mg/ml ethidium bromide to be separated on a 1% formaldehyde agarose gel in 1 x MOPS (20 mM 3-[N-morpholino]propanesulfonic acid, 5 mM sodium acetat, 1 mM EDTA, final pH 7.0) running buffer. After electrophoresis, RNAs were blotted onto a Hybond-NX membrane (GE Healthcare) overnight by capillary transfer using 5 x SSC buffer (0.75 M NaCl, 0.075 M sodium citrate). After disassembly of the capillary transfer apparatus, the membrane was rinsed in 0.5 M NaOH followed by extensive wash with 5 x SSC and fixed with 1200 J UV. The membrane was incubated in blocking solution (10% SDS, 1 mM EDTA in PBS) for 20 min and probed with 1 µg/ml streptavidin-HRP in blocking solution for 15 min. The membrane was then washed six times in PBS containing decreasing concentrations of SDS (10%, 1%, 0.1% SDS, twice each and for 10 min). The signal was visualized using ECL detection reagents (Thermo). Detection of 4sU-labeled RNA by RT-qPCR (II) After biotinylation and precipitation, 30 µg RNA was diluted in 100 µl of RNAse-free water. The 4sU-labeled RNA was captured using µMACS streptavidin beads and columns (Miltenyi). Briefly, 100 µl beads were transferred and magnetically fixed to each column followed by 3 washes with washing buffer (100 mM Tris pH 7.5, 10 mM EDTA, 1 M NaCl). The beads were then recovered in 100 µl washing buffer by removing the column from the magnetic area and incubated with RNA for 15 min with rotation. The 50 reaction volume was applied to µMACS columns placing in magnetic stand and washed twice with 300 µl of 65 °C washing buffer followed by 4 washes with room temperature washing buffer. The beads were collected in 100 µl of cDNA reactions with TaqMan MULV reverse transcriptase and random hexamer (Applied Biosystems) for cDNA synthesis, after which the mixture was applied to µMACS columns and the cDNA was collected as flow-through followed by RT-qPCR analysis as described. Statistical analyses (I-IV) Statistical analyses were performed using t test calculator from GraphPad QuickCalcs (http://www.graphpad.com/quickcalcs/ttest1/; January, 2014). 51 RESULTS 1. Cdk7-cyclin H-Mat1 kinase subcomplex of TFIIH is required for global transcription and mRNA turnover (I, II) The contradictory evidences from different models regarding the in vivo Pol II CTD Ser5 phosphorylating kinase and the role of the TFIIH-associated kinase subunit in general transcription in mammalian cells prompted us to investigate the global transcription effects following depletion of either Cdk7 or Mat1. Mat1 was depleted in mouse embryonic fibroblasts (MEFs) carrying a floxed Mat1 allele through adenoviral infection expressing Cre recombinase (Korsisaari et al., 2002). At 72 hours post-infection, Mat1 and Cdk7 proteins were undetectable by western blot and the cellular level of Ser5 phosphorylation of Pol II CTD was significantly decreased as demonstrated by western blot and immunostaining (I). Chromatin-associated Ser5P level was also decreased to 1025% in the four analyzed genes as assayed by chromatin immunoprecipitation (ChIP) (I). These results demonstrate that Mat1 is required for Ser5 phosphorylation of Pol II CTD and suggest that Cdk7 is the major Ser5 phosphorylating kinase in mammalian cells. Intriguingly, downregulation of Cdk7 using siRNA targeting murine Cdk7 in immortalized MEFs did not result in global decrease in Ser5P (II) consistent with reports on Cdk7-depleted MEFs (Ganuza et al., 2012) and human colon cancer cells following chemical inhibition of Cdk7 activity (Glover-Cutter et al., 2009; Larochelle et al., 2012). However, ChIP analysis revealed a significant decrease in Ser5P in gene promoters by acute reduction of Cdk7 at 48 h following siRNA transfection in MEFs (II). Unexpectedly, gene expression profiling of steady-state mRNAs did not reveal global changes in Mat1-depleted MEFs with impaired Ser5 phosphorylation. Less than 1% of mRNA levels were altered in Mat1-depleted MEFs by conventional criteria (I). ChIP analyses demonstrated decreased Pol II levels in the bodies of all genes examined, including genes without any changes in abundance (I). This suggested a widespread defect on Pol II elongation that was not reflected by the steady-state mRNA levels in Mat1-depleted cells. To analyze the amount of newly synthesized mRNAs instead of the steady-state levels, 52 Mat1-delpeted MEFs were subjected to transcriptional induction by serum stimulation or heat shock. Measurements of c-Fos and Hsp70 mRNAs upon induction revealed a defective and delayed accumulation of these two genes following Mat1 depletion (I). A similar defective induction of Hsp70 was also observed in Cdk7 knockdown MEFs together with a significant decrease in Ser5P at the promoter (II), consistent with a previous report in Drosophila (Schwartz et al., 2003). This prompted us to analyze the global transcription by metabolic pulse labeling of nascent RNA using the ribonucleoside analog 5-fluorouridine (FUrd) and 5-ethynyl uridine (EU) followed by 1 h chase. This type of analyses revealed a significant decrease in the total amount of labeled RNAs in Mat1-depleted MEFs (I). Similar results were obtained using a short pulse of FUrd in addition to 4-thiouridine (4sU) in Cdk7 knockdown MEFs (II). A decrease in newly synthesized RNAs was also observed in human HEK293 cells following transfection of siRNA targeting endogenous human CDK7, which can be rescued by expression of wildtype murine Cdk7 (Cdk7-WT), but not the kinase-deficient (Cdk7-K33M), demonstrating that the kinase activity of Cdk7 is required for general transcription (II). Analysis of 4sU-incorporated RNAs using gene-specific primers also revealed a decrease in the synthesis of 18S and 28S rRNA but not the Pol III-transcribed gene 7SL in Cdk7deregulated cells (II). This is consistent with previous reports indicating an involvement of TFIIH in Pol I-mediated transcription of rDNA (Bradsher et al., 2002; Hoogstraten et al., 2002; Iben et al., 2002). Interestingly, the decreased 4sU incorporation was correlated with reduced H3K4trime associated with both the Pol II and Pol I genes (II), also indicating an attenuation of transcription of these genes. The effect on rRNAs by Cdk7 knockdown is not secondary to the inhibition of Pol II transcription, as treatment of MEFs with low doses of α-amanitin for 48 h only led to a negligible decrease in 18S and 28S (II). Three mRNAs without changes in the steady-state level were chased for more time points till 72 h and displayed a much longer lifetime after Mat1 depletion (I), suggesting that the transcription attenuation in these cells might be compensated by increased mRNA stability. This conclusion is supported by a general stabilization of labeled RNAs in Cdk7 knockdown MEFs shown by pulse-chase of the labeled RNAs (II). 53 In addition to the global transcriptional attenuation, a few specific mRNAs were found associated with capping and splicing defects following Mat1 depletion (I), consistent with current knowledge of CTD phosphorylation in coupling transcription and cotranscriptional mRNA processing. Taken together, the study on Mat1 and Cdk7 demonstrated that mammalian Cdk7 kinase subcomplex is required for global transcription and mRNA turnover. The finding that the defects in transcription are not propagated to changes in the steady-state levels in some cases reveals the limitation in previous mammalian studies (Ganuza et al., 2012; GloverCutter et al., 2009; Larochelle et al., 2012) in regard of Cdk7 in transcriptional regulation. 2. Cdk7 and the associated Mat1 inhibits adipogenic transcription program through inhibitory phosphorylation of PPARγ (III) To study the potential cell type-specific functions of TFIIH kinase submodule, the physiologic levels of Cdk7 and Mat1 in different tissues were examined. In contrast to the presumably ubiquitous expression, Cdk7 and Mat1 proteins were undetectable in terminally differentiated white adipocytes in mice. The reduced Cdk7 and Mat1 levels noted in adipocyte tissues were recapitulated in the 3T3-L1 adipocyte differentiation model, where both subunits decreased progressively during the conversion of preadipogenic fibroblasts into adipocytes induced by hormones. To explore the functional relevance of this modulation of Cdk7 and Mat1 levels for adipogenesis, Mat1-depleted MEFs were induced to adipogenic differentiation by hormone treatment. Interestingly, these cells displayed an enhanced ability of differentiating into adipocytes, which could be reversed by Mat1 restoration, suggesting that the Cdk7 subcomplex inhibits adipogenesis. Because of the critical role of PPARγ in adipogenesis (Rosen et al., 1999; Tontonoz et al., 1994) and the tentative link to Cdk7 (Compe et al., 2005), the level of PPARγ S112 phosphorylation was examined in U2OS cells following transfection of plasmid coding for GFP, PPARγ and shRNA targeting Cdk7 (shCdk7) or non-targeting (shControl). In transfected cells, as indicated by GFP expression, the Cdk7 staining was undetectable in shCdk7 cells comparing to shControl cells, as well as the staining of S112 specific phosphorylated PPARγ, but the signal for 54 total PPARγ was intact. This demonstrates that Cdk7 is required for the S112 phosphorylation of PPARγ in culture cells, consistent with the reported in vitro kinase activity of Cdk7 towards this site (Compe et al., 2005). An increased PPARγ-mediated transcriptional response was noted by examining the PPARγ reporter activity and the endogenous levels of two PPARγ targeted genes (adiponectin and aP2) in MEFs depleted of Cdk7 or Mat1. This acquired responsiveness is dependent on PPARγ S112 because the increased reporter activity following Cdk7 knockdown was eliminated when S112 was mutated to a non-phosphorylatable alanine. These results together demonstrated that the Cdk7 submodule of TFIIH inhibits PPARγ-mediated adipogenic transcription through phosphorylating PPARγ S112 and acts as a physiological roadblock to adipogenesis. 3. Inhibition of ciliogenesis by CCRK and its effectors ICK and MAK promotes cell proliferation (IV) To study whether the ciliary function of CCRK is evolutionally conserved in mammals, CCRK level was reduced by siRNA in NIH3T3 mouse fibroblasts followed by serum starvation to promote ciliogenesis. An 80% reduction in CCRK mRNA level resulted in more ciliated cells at 8 h of serum starvation and attenuated cilia disassembly induced by serum restimulation, indicating that CCRK downregulation promotes cilia formation and increases cilia stability. After serum starved for 24 h, almost all NIH3T3 cells were ciliated and the ciliation was not significantly altered despite that the cilia were longer in CCRK knockdown cells. The elongation of cilia was rescued by expression of siRNAresistant wild-type (WT) CCRK but not the kinase-deficient CCRK. These results established that mammalian CCRK negatively regulates ciliogenesis through its kinase activity. No cilia curling was observed in NIH3T3 cells following CCRK knockdown, in contrast to a report on motile cilia in CCRK-deficient zebrafish kidney (Ko et al., 2010), suggesting that CCRK regulation of ciliogenesis is context and possibly dose dependent as has been shown for LF2 in regulating Chlamydomonas flagella (Tam et al., 2007). It is of interest to note that knockdown of Cdk7 did not elongate cilia (my unpublished data). The elongated cilia in CCRK-depleted cells were associated with elevated Hedgehog (Hh) transcriptional responsiveness as demonstrated by increases in the Gli-luciferase reporter 55 activity and endogenous mRNA levels of two Hh target genes (Gli1 and Ptch) upon 4048 h treatment of ligand. Immunostaining of Gli3 revealed more cells with Gli3 at ciliary tips following CCRK knockdown 1 h after ligand stimulation, suggesting that CCRK knockdown leads to increased activation of an early event in Hh signaling in conjunction with increased downstream transcriptional responses. Despite the full competence to rescue, overexpressed WT CCRK was not enriched in cilia or basal bodies, suggesting an indirect role of CCRK in regulating ciliogenesis. Overexpression of ICK, a kinase previously reported to be phosphorylated by CCRK, reduced ciliation dramatically (from 88.8±3.0% in vector control to 11.1±3.5%). Interestingly, in the rare ciliated cells expressing exogenous ICK, ICK was found enriched in the distal tips of cilia. Consistent with an inhibitory role of ICK in regulating ciliogenesis, knockdown of ICK in NIH3T3 cells led to cilia elongation comparable to CCRK knockdown. Interestingly, the inhibition of cilia formation in ICK overexpressed cells was severely blunted following CCRK knockdown (ciliation from 11.1±3.5% to 59.4±6.2%), demonstrating that CCRK is required for ICK-mediated inhibition of ciliogenesis. Cells expressing ICK with reduced CCRK level demonstrated prominent cilia, and again ICK was enriched at distal tips. ICK mutated at the CCRK phosphorylation site to a non-phosphorylatable alanine (T157A) also localized to the ciliary tip but displayed a severely blunted ability to suppress cilia formation compared to ICK-WT (T157A, 55.8±4.2%; WT, 11.1±3.5%). The remaining suppression (compared to 88.8±3.0% in vector control) is likely due to the residual activity of ICK-T157A (Togawa et al., 2000). In addition, overexpression of an ICK phosphomimetic mutant (T157E) reduced cilia frequency comparably to ICK-WT (T157E, 8.3±7.6%; WT, 11.1±3.5%) and ICK-T157E in rare ciliated cells was also enriched at the ciliary tips. In contrast to ICK-WT, the ciliation in ICK-T157E cells was not significantly changed following CCRK knockdown. These data together indicated that the ciliary function of CCRK is mediated through ICK-T157 phosphorylation. Downregulation of CCRK and ICK levels caused an inhibition of G1/S transition revealed by qualification of replicating cells using EdU labeling in NIH3T3 cells upon serum restimulation. The cell cycle effect is dependent on cilia because knockdown of 56 CCRK or ICK did not change the percentage of S phase cells in cells stably expressing shRNA targeting Kif3a, an IFT motor subunit required for maintaining cilia (Marszalek et al., 1999). In Kif3a-depleted cells, ciliogenesis was suppressed as evidenced by an increase in the percentage of cells with absent or short cilia. Importantly, knockdown of CCRK or ICK did not increase the length of cilia in Kif3a-depleted cells. As cilia have been implicated in proliferation in the context of cancer (Han et al., 2009) (Wong et al., 2009) and CCRK has been identified as an overexpressed oncogene promoting cell proliferation in glioblastoma (Ng et al., 2007), I next investigated whether the loss of cilia in glioblastoma cells is due to CCRK overexpression and whether this is associated with increased proliferation. In glioblastoma cells with deregulated high levels of CCRK, reducing CCRK level by siRNA restored cilia in a subset of cells that were significantly deficient in S-phase entry as shown by co-staining of cilia and EdU. In these cells an activated PI3K pathway contributes to deregulated CCRK expression and the downstream effectors of CCRK in suppressing ciliogenesis include ICK together with an ICK-related kinase MAK (male germ cell-associated kinase), which is not expressed in NIH3T3 cells. Importantly, CCRK reduction led to a proliferation defect in these cells largely dependent on cilia because the defect was significantly rescued by depletion of Kif3a and Ift20 to prevent cilia formation. Together, these results demonstrated that the inhibition of ciliogenesis by overexpressed CCRK promotes proliferation of glioblastoma cells. 57 DISCUSSION & PERSPECTIVES 1. Cdk7 subcomplex of TFIIH in transcription mediated by RNA Pol II and Pol I The study conducted here by analyzing nascent RNA reveals a requirement of Cdk7 submodule for general transcription in mammalian cells. Our work also demonstrates that the function of Cdk7 in general transcription is dependent on its kinase activity. The decreased levels of newly synthesized mRNAs by Pol II was associated with a reduction of CTD Ser5P level on the chromatin as well as a hallmark of actively transcribed chromatin (H3K4me3) of which the catalytic enzyme has been reported to be recruited by Ser5-phosphorylated CTD (Ng et al., 2003). At this point, we could not distinguish whether the remaining Ser5P is due to the residue level of Cdk7 in our system or the existence of additional CTD Ser5 phosphorylating kinases as suggested by previous studies (Ganuza et al., 2012; Glover-Cutter et al., 2009; Larochelle et al., 2012; Rossi et al., 2001). Size separation and localization analysis of the nascent RNAs suggests that the synthesis of rRNA mediated by Pol I is also defective in Cdk7-deficient cells, which is confirmed by using specific primers for rRNAs (II). These results demonstrate that the mammalian Cdk7 is required for Pol I-mediated rDNA transcription. Inherited mutations in the XPB subunit of TFIIH reduce the binding of Cdk7 to rRNA genes, accompanied by an impaired rRNA synthesis (Assfalg et al., 2012). As Pol I has intrinsic ATP hydrolysis activity and does not require XPB to hydrolyze ATP (Vannini and Cramer, 2012), the function of XPB in Pol I transcription might be dependent on Cdk7. The activities of Pol I and Pol II are substantially different and current in vitro biochemistry data does not support a similar role of TFIIH in Pol I transcription as in Pol II regulating at the initiation step (Assfalg et al., 2012; Iben et al., 2002). Interestingly, many of the CTD kinases and a phosphatase Fcp1 also genetically interact with the universally conserved transcription elongation factor Spt5 and its binding partner Spt4 (Hartzog and Fu, 2013; Lindstrom and Hartzog, 2001). Moreover, many of the CTD kinases including Cdk7 homolog Kin28 have been implicated in phosphorylating the Cterminal region (CTR) of Spt5 (Liu et al., 2009; Qiu et al., 2012; Yamada et al., 2006), 58 which appears to act as a phosphorylation-state dependent regulator recruiting transcription-associated chromatin modifiers, resembling the Pol II CTD (Hartzog and Fu, 2013). It would be interesting to test whether the mammalian Cdk7 is involved in the phosphorylation of Spt5 CTR and in turn the potential implications on the elongation step of transcription mediated by Pol II and Pol I. 2. Crosstalk between RNA synthesis and degradation: an emerging layer of regulation for gene expression The defects in nascent RNA synthesis do not lead to decreased steady-state levels in Cdk7 or Mat1-deficient cells. We proposed that this uncoupling is due to a compensative increase in the RNA stability (Figure 3) as the pulse-labeled RNAs were resistant to degradation after hours of chase both in Mat1-deleted cells and Cdk7-deficient cells (I and II). This indicates that the primary changes in transcription could be masked by assays measuring the steady-states level only, providing a rationale for the failure of identifying the general requirement of Cdk7 for Pol II and Pol I transcription in the previous studies (Ganuza et al., 2012; Glover-Cutter et al., 2009; Larochelle et al., 2012). Our study also demonstrates that, as in yeast, a widespread RNA buffering mechanism also exists in mammals, representing a previously overlooked layer of regulation for gene expression. The pioneering studies in yeast have identified two Pol II subunits Rpb4 and Rpb7 coupling the transcription in the nucleus and mRNA decay in the cytoplasm (DoriBachash et al., 2011; Goler-Baron et al., 2008). Interestingly, the Rpb4/Rpb7 heterodimer locates in the vicinity of the CTD of Rpb1 in the complete enzyme (Bushnell and Kornberg, 2003). It would be interesting to study whether Rpb4/7 is involved in the increased mRNA stability in Cdk7 or Mat1-deficient cells, associated with decreased Ser5 phosphorylation of CTD (Figure 3). 59 wild-type cells Cdk7-deficient cells Nucleus Nucleus mRNA decay factor imprinting factor ribosome amino acid Figure 3. Simplified scheme of proposed compensation of decreased transcription by increased mRNA stability in Cdk7-deficient cells comparing to the wild-type cells. In the wild-type cells, Cdk7 phosphorylates Pol II CTD and activates Pol II-mediated transcription to produce mature mRNAs associated with proper imprinting factors (such as Rbp4/7) instructive to the decay factors mediating mRNA degradation. When Cdk7 is depleted, Pol II is hypophosphorylated and less effective in transcription. The recruitment of the imprinting factors to promoters is also defective due to loss of Cdk7 or Pol II CTD phosphorylation, resulting in mRNAs without imprinting factors. The half-lives of these mRNAs are increased because the decay factors are unable to recognize them efficiently without imprinting factors. K7, Cdk7; P, phosphorylated. 3. Cell type-specific functions of Cdk7 subcomplex The link between Cdk7 and PPARγ in vivo has been suggested previously by a study in mice with mutations in the XPD subunit of TFIIH (Compe et al., 2005). The genetically modified XPD caused deregulation of several PPARγ target genes in adipose tissue and was proposed to affect Cdk7 activity towards PPARγ Ser112 (Compe et al., 2005). Our results demonstrate that Cdk7 is required for phosphorylation of PPARγ Ser 112 and represses the expression of PPARγ target genes through this inhibitory phosphorylation 60 (III). Our findings that Cdk7 and Mat1 are undetectable in the murine adipose tissue and decreased during adipogenesis of 3T3L1 preadipocytes (III) demonstrate that the Cdk7 subcomplex is a physiological roadblock of adipocyte differentiation. Moreover, it is important to notice that MEFs from the PPARγ S112A mutant mice display a similar enhanced adipogenic potential (Rangwala et al., 2003) as Mat1-depleted MEFs, which further supports that the Cdk7 subcomplex inhibits adipogenesis by phosphorylating PPARγ in vivo. The absence of the Cdk7 subcomplex subunits of TFIIH in adipose indicates that the function of TFIIH is tissue specific. It would be interesting to study the Ser5P level of Pol II CTD in adipocytes as well as the relevance to basal transcription. It is also of great interest to examine the expression level of other TFIIH subunits as well as their involvement in the basal transcription. It is possible that TFIIH is missing in the adipose tissue as a holocomplex or it adapts its subunit composition to the adipogenic program. For almost 30 years, it was thought that the core transcription machinery is an invariant and universal entity in eukaryotic cells. Combining with the observation that the canonical TFIID subunits are replaced during the differentiation of muscle cells (Deato and Tjian, 2007), this concept has been overturned and the core transcription machinery in differentiated cells seems to have very diverse compositions. 4. The role of CCRK and its substrates ICK/MAK in regulating ciliogenesis Our studies on mammalian CCRK reveal a conserved role of CCRK in regulating ciliogenesis and suggest that this regulation is mediated through phosphorylation of two related kinases ICK and MAK at their TDY motifs. This study provides the first functional relevance for this phosphorylation event. The enormous progresses in the past decade in understanding the formation and functions of primary cilia, especially the findings of “ciliopathies” in humans have made this field of great medical importance. The research efforts devoted to unravel the physiological ways regulating ciliogenesis should be encouraged. The effectors of CCRK- ICK and MAK are especially interesting because of their extraordinary localization at a barely studied area- the ciliary tips (III and Omori et al., 2010) where IFT particles switch from 61 anterograde to retrograde transport and microtubules are constantly turning over (Ishikawa and Marshall, 2011). Identifying mediators for the ciliary function of ICK/MAK may help reveal the molecular mechanisms of these two processes as well. Two microtubule-associated ciliary protein, Retinitis pigmentosa 1 (RP1) and Rp1-like protein, have been indicated to mediate the cilia elongation in MAK-depleted mouse photoreceptor cells (Omori et al., 2010). The exclusive expression of RP1 and Rp1-like protein in photoreceptor cells (Yamashita et al., 2009) indicates that they do not mediate the effects of ICK/MAK on ciliogenesis in fibroblasts or glioblastoma cells observed here. The observed increase in ciliary translocation of Gli3 (an IFT cargo) associated with CCRK downregulation (IV), together with reported abnormal accumulation of certain IFT components in CCRK/ICK/MAK homologue mutants and in Mak-null mouse photoreceptor cells (Burghoorn et al., 2007; Omori et al., 2010; Phirke et al., 2011). (Burghoorn et al., 2007; Phirke et al., 2011) suggests that CCRK and ICK/MAK inhibit ciliogenesis through regulating IFT. Knockdown of CCRK or ICK does not elongate the remaining cilia in cells with reduced Kif3a (IV) further suggesting that the ciliogenesis regulation by CCRK and ICK is IFT dependent. Our results also suggest a functional redundancy of MAK with ICK, providing a rationale for lack of ciliary defects in Mak-null mice in most of tissues (Shinkai et al., 2002). It would be interesting to study the expression patterns of these two kinases for potential tissue-specific functions. 5. Cilia, cell cycle and cancer This study has linked the inhibitory ciliary function of CCRK and ICK to S-phase entry, together with previous publications (Kim et al., 2011b; Li et al., 2011), suggesting that primary cilia may act as a physical checkpoint for G1/S transition (Figure 4). Future studies on the molecular basis for this inhibition of the cell cycle posed by cilia have immediate implications for pathogenesis dependent on the transition from quiescence to a proliferative state, such as cancer, polycystic kidney and neocortical development. Our studies in glioblastoma cells indicate that CCRK overexpression is a mechanism by 62 which glioblastoma cells deregulate a pathway operational in normal cells to eliminate cilia and promote growth. Downregulation of CCRK or its effectors is able to restore cilia in glioblastoma cells resulting in a block to S-phase entry (IV). Interestingly, the cilia restoration could also be achieved by a small molecule reducing the CCRK mRNA levels (IV). Therefor, to develop small molecules specifically targeting ciliogenesis regulators might provide novel strategies for treatment of glioblastoma. low CCRK ICK$ high CCRK Anterograde P$ ICK$ P$ICK$ ICK$ Retrograde ICK$ ICK$ ICK$ P$ICK$ P$ICK$ P$ICK$ P$ICK$ P$ P$ICK$ ICK$ ICK$ ICK$ ICK$ P$ICK$ ICK$ P$ICK$ P$ICK$ P$ICK$ Figure 4. Schematic diagrams depicting the proposed model of CCRK and its substrate kinase ICK regulating ciliogenesis and cell cycle progression. In the left, the phosphorylated and activated ICK by CCRK is transported into primary cilium through IFT, where it inhibits the anterograde transport or/and promotes the retrograde transport. The unphosphorylated ICK is inactive and accumulates in the tip. The competition for the anterograde IFT complex to enter cilium between the phosphorylated and unphosphorylated ICK limits the ICK activity to a proper level, resulting in the steady-state ciliary length. The presence of cilium inhibits the G1/S transition of the cells cycle. In the right, the high level of CCRK (for example in glioblastoma cells) leads to more phosphorylated ICK and the enhanced ICK activity results in cilium disassembly, followed by loss of the inhibitory effect of cilium on cell cycle progression. P, phosphorylated. 63 CONCLUSIONS This study aimed at comparing functions of the closely related Cdk7 and CCRK kinases has identified distinct cellular functions of Cdk7 and CCRK. Nascent RNA analysis reveals a general requirement of the kinase activity of Cdk7 for both Pol II and Pol I-driven transcription in mammalian cells. The defective transcription following disruption of the Cdk7 submodule is accompanied with a compensative increase in RNA stability, identifying a coupled regulation mechanism between transcription and RNA decay in mammalian cells. The results on the Cdk7 submodule of TFIIH in adipogenic program reveal cell typespecific functions of the Cdk7 subcomplex subunits and demonstrate that the Cdk7 subcomplex inhibits adipocyte differentiation involving Cdk7-mediated inhibitory phosphorylation of PPARγ. This study therefore contributes the understanding of adipogenesis mediated by PPARγ and links modulation of basal transcription machinery to differentiation. The study on CCRK identifies an important function in primary cilia regulation and reveals a CCRK-ICK/MAK kinase cascade that inhibits ciliogenesis in mammalian cells. The results also demonstrate that this pathway is modulated in glioblastoma cells with high CCRK expression level to eliminate primary cilia thereby promoting proliferation. The causal link established here between cilia and proliferation together with the identified potential of cancer cells to restore cilia should stimulate exploring cilia restoration as a therapeutic opportunity in cancer. 64 ACKNOWLEDGEMENTS This study was carried out at Biomedicum Helsinki and Institute of Biotechnology, University of Helsinki during 2006-2014 under the supervision of Professor Tomi Mäkelä. I thank the staff of Biomedicum Helsinki and Institute of Biotechnology for excellent research facilities, especially Eija Koivunen and Mervi Lindman. The Helsinki Graduate Program in Biotechnology and Molecular Biology (GPBM) is acknowledged for years of financial support and a great set of courses. Special thanks to GPBM coordinator Dr. Erkki Raulo and Professor Deyin Guo at Wuhan University for initiating the summer school program where I got to know the scientific community in Finland and found an opportunity to do a PhD at University of Helsinki. This study has also been financially supported by Biocentrum Helsinki, Finnish Cancer Organization, Sigrid Juselius Foundation, Academy of Finland, University of Helsinki, Finnish Cultural Foundation, Instrumentariumin Tiedesäätiö, Ida Montinin Säätiön, Biomedicum Helsinki Foundation, the Company of Biologists Limited and Magnus Ehrnrooth foundation. I am extremely grateful to my supervisor professor Tomi Mäkelä who has provided me with an excellent lab with an inspiring atmosphere for both science and fun. I really appreciate your guidance from my baby steps in the lab as an undergraduate student from China to much more intensive discussions at the later stage. Your “razor sharp wits” are behind all the scientific progress that I may have achieved. Thank you especially for your unreserved support when I chose to explore a “non-lab-main-stream” topic. Thank you also for your trust, because of which you have allowed me to conduct my study in my own way and I will always appreciate it! I thank Lea Sistonen and Päivi Ojala for being my thesis committee follow-up members. Thank you so much for your valuable inputs on my research projects as well as my other studies. The annual meetings we have had were really serving as “check-points” and I feel fortunate that I have had the chance to discuss with you every aspect throughout my PhD study. I thank Lea Sistonen (again) and Markku Varjosalo for reviewing my thesis and providing me with numerous valuable comments that further strengthened the thesis. I thank all my co-authors without whom this work would not be possible. Particular thanks go to Katja Helenius, who initiated the studies on the transcriptional functions of TFIIH and has always been a role model since the beginning when I started in the Mäkelä lab. Your enthusiasm towards science and highly efficient work style will continue to inspire me and I deeply appreciate the opportunity of working with you on these projects. 65 I want to devote my warmest gratitude to both past and present members of the Mäkelä lab. I am just really lucky to be with you during these years! In particular I want to thank Niina Roine, Heini Hakala, Saana Ruusulampi, Daria Blokhina and Timofei Tselykh for superb hands-on help and providing data with significant impact on our studies. Our past and present technicians- Saana Ruusulampi (again), Niina Roine (again), Markus Keskitalo, Kaisa Laajanen, Outi Heikkilä, Susanna Räsänen, Kirsi Mänttäri, Sari Räsänen and Jenny Bärlund- thank you so much for your excellent work to keep the lab running smoothly. Special thanks go to Niina (again), Saana (again) and Markus (again) for understanding my particular (or picky) requirements for reagents and always making sure I get what I want in time. I also want to thank the “transcription people in lab”- Thomas Westerling, Katja Helenius (again), Emilia Kuuluvainen, Timofei Tselykh (again) and Heini Hakala (again) for fruitful scientific discussions and terrific company in the office/at the bench. In addition, I thank all the Mäkelä lab members for your scientific inputs and countless occasions we spent together for fun. Special thanks to Lina, Jing, Yan, Iris and Jessica for friendship and sharing interests in cooking. The Frilander lab, Klefalab and Ojalalab are also acknowledged for exchanging ideas and sharing reagents as well as the leisure times during lab retreats/lunch breaks. I also thank Satu Sankkila, Jonna Katajisto, Anita Tienhaara, Aija Kaitera and Anu Taulio for administrative help without which I wouldn’t have been able to meet the deadlines in many cases. Thanks to my other Chinese friends in Finland: Fang Chen, You Zhou, Ping Chen, Yizhou Hu, Songping Li, Miao Yin, Shengtong Fang, Li Ma, Hongqiang Ma, Xiaonan Liu, Li Tian, Chunmei Li, Hong Wei, Bei Yang, Ying Liu, Jianping Chen, Qiang Lan and Jing Cheng for your advices and encouragements that have helped me in so many ways and surely made me less homesick. Thanks to my parents and parents-in-law for your endless support. Thanks to Mom and Dad for raising me in such a way that you always encourage me to make my own choices and pursue my dream even though this means that we have to live in two continents for quite a long time. Finally, my love Miaoqing, thank you so much for your unconditional support, thank you for believing in me that I am capable of doing what I want to do, thank you for reminding me all the time how much I like doing science, thank you for understanding my schedule, thank you for sharing my ups and downs… This is going to be an endless list and all in one- thank you for being in my life and with me for this long journey. Helsinki, May 22nd, 2014 66 REFERENCES Adams, M., Reginato, M.J., Shao, D., Lazar, M.A., and Chatterjee, V.K. (1997). Transcriptional activation by peroxisome proliferator-activated receptor gamma is inhibited by phosphorylation at a consensus mitogen-activated protein kinase site. J Biol Chem 272, 5128-5132. Adelman, K., and Lis, J. (2012). Promoter-proximal pausing of RNA polymerase II: emerging roles in metazoans. Nature reviews Genetics 13, 720-731. Akhtar, M.S., Heidemann, M., Tietjen, J.R., Zhang, D.W., Chapman, R.D., Eick, D., and Ansari, A.Z. (2009). 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