The Stabilization of Basal Bodies to Resist Cilia Generated Forces

Home , Axoneme, Basal body, Nexin

THE STABILIZATION OF BASAL BODIES TO RESIST CILIA GENERATED FORCES

BRIAN ANTHONY BAYLESS

B.S., University of California, Irvine, 2010

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Cell Biology, Stem Cells, and Development

2016

This thesis for the Doctor of Philosophy degree by

Brian Anthony Bayless

has been approved for the

Cell Biology, Stem Cells, and Development Program

Michael McMurray, Chair

Bruce Appel

Jennifer DeLuca

Jeff Moore

Rytis Prekeris

Chad Pearson, Advisor

Date: August 19, 2016

Bayless, Brian Anthony (Ph.D., Cell Biology, Stem Cells and Development)

The Stabilization of Basal Bodies to Resist Cilia Generated Force

Thesis directed by Assistant Professor Chad G. Pearson

ABSTRACT

Centrioles and basal bodies (CBBs) are the major microtubule-organizing centers of the cell. Centrioles nucleate the microtubules of the mitotic spindle, and basal bodies anchor and nucleate the ciliary/flagellar axoneme. In both contexts, centrioles and basal bodies experience mechanical force from the pulling of chromosomes during mitosis and the movement of the cilia and flagella, respectively. Failure to stabilize against mechanical force causes CBBs to fragment or disassemble, and may lead to multipolar mitosis and disassembly of cilia. Structurally, CBBs are radially symmetric cylinders comprised of nine sets of modified triplet microtubules arranged around a cartwheel shaped structure at its proximal end. The triplet microtubules that make up CBBs are hyperstable as evidenced by their resistance to various microtubule stressors. The factors that stabilize the CBB from mechanical force are unknown. The ciliate Tetrahymena thermophila serves as a promising model organism to study mechanical forces on basal bodies. Using this system I have shown that basal body stability proteins Bld10 and Fop1 stabilize basal bodies against the forces of ciliary beating. I have also developed a novel method to measure the incorporation dynamics of proteins into the basal body. Additionally, I have shown that Fop1 and post- tubulin glutamylation localize asymmetrically at the basal body coincident with the site of highest predicted cilia generated compression forces. This work highlights my achievements and contributions to the understanding of CBB stabilization against mechanical forces.

The form and content of this abstract are approved. I recommend its publication.

Approved: Chad G. Pearson

iii

TABLE OF CONTENTS

CHAPTER

I: INTRODUCTION ...... 1

Centrioles and Basal Bodies ...... 1

Centriole and Basal Body Stabilization ...... 11

Motile Cilia and Flagellar Beating ...... 20

Tetrahymena as a Model Organism for Basal Body Research ...... 30

Conclusions and Thesis Aims ...... 37

II: BLD10/CEP135 STABILIZES BASAL BODIES AGAINST CILIA-GENERATED FORCE.39

Introduction ...... 39

Results ...... 42

Discussion ...... 62

Materials and Methods ...... 67

III: MOLECULAR ASYMMETRIES STABILIZE BASAL BODIES AGAINST CILIARY BEATING FORCES...... 74

Introduction ...... 74

Results ...... 77

Discussion ...... 96

Materials and Methods ...... 99

IV: CONCLUSIONS AND FUTURE DIRECTIONS ...... 105

APPENDIX A...... 110

APPENDIX B...... 113

APPENDIX C ...... 117

REFERENCES ...... 118

LIST OF FIGURES

1.1: CBB structure ...... 5

1.2: Axoneme structure ...... 22

1.3: Polarized organization of Tetrahymena basal bodies ...... 31

1.4: Tetrahymena basal body structure ...... 32

1.5: Schematic representation of Tetrahymena basal bodies and associated accessory structures ...... 34

2.1: TtBld10 is a member of a highly conserved family of CBB proteins ...... 43

2.2: TtBld10 localizes to the basal body outer cartwheel domain ...... 45

2.3: TtBld10 loss causes decreased cellular growth ...... 48

2.4: TtBld10 knockout causes a loss of basal bodies...... 49

2.5: Maturation and disassembly of K-like-Antigen ...... 50

2.6: TtBld10 is required for basal body assembly ...... 51

2.7: TtBld10 is required for the maintenance of basal bodies ...... 53

2.8: Triplet microtubule assembly and stability defects in TtBld10 cells ...... 55

2.9: TtBld10 protein stably accumulates at basal bodies ...... 58

2.10: GFP-TtBld10 temporally matures at basal bodies ...... 60

2.11: Cilia generated forces destabilize basal bodies in TtBld10 cells ...... 61

3.1: Fop1 is a basal body stability protein ...... 78

3.2: Supplemental to Figure 3.1 ...... 79

3.3: Fop1 stabilizes basal bodies to resist ciliary beating ...... 81

3.4: Fop1 localizes asymmetrically at the basal body ...... 83

3.5: Supplemental to Figure 3.4 ...... 86

3.6: Poc1 promotes normal incorporation of Fop1 into the basal body ...... 87

3.7: Supplemental to Figure 3.6 ...... 89

3.8: Basal body microtubule glutamylation increases in basal body stability mutants .... 90

3.9: Supplemental to Figure 3.8 ...... 93

3.10: Basal bodies are stabilized through distinct pathways ...... 94

3.11: Supplemental to Figure 3.10 ...... 95

ABBREVIATIONS

CBB: Centriole and Basal Body

PCM: Pericentriolar Material

MAP: Microtubule Associated Protein

IEM: Immuno-Electron Microsopy

Kl-Ag: K-like-Antigen

TEM: Transmission Electron Microscopy

FRAP: Fluorescent Recovery After Photobleaching

PEO: Poly Ethelene Oxide

PTM: Post-Translational Modification

SIM: Structured Illumination Microscopy

CR: Conserved region

vii

CHAPTER I

INTRODUCTION1

Centrioles and Basal Bodies

Centrioles and basal bodies (CBBs) are self-assembling cellular structures that act as microtubule organizing centers. As microtubule organizing centers, CBBs are responsible for organizing the microtubules of mitotic spindles and ciliary axonemes, respectively. CBBs are composed of modified triplet blades of microtubules that extend to make up the length of their barrel shaped structure. The integrity of this structure is essential for centrioles and basal bodies to serve their function as microtubule organizing centers. In order to ensure proper structural integrity of centrioles and basal bodies, their assembly and maturation is highly regulated and organized.

Centriole and basal body function

CBBs are evolutionarily conserved microtubule organizing centers. During mitosis centrioles are key components of centrosomes, which nucleate the microtubules of the mitotic spindle. As the cell enters G0 or G1 phase of the cell cycle, the centriole can migrate to the cell cortex and nucleate the microtubules that comprise the ciliary axoneme. At this point the centriole is referred to as a basal body. CBBs are found in most eukaryotes including animals, early plants, and flagellates (Marshall, 2009; Yubuki and Leander, 2013).

Their existence is strongly correlated with whether that species evolved a cilium or flagella, suggesting that microtubule organizing centers first evolved as basal bodies and only later gained functionality as centrioles (Marshall, 2009). Organization of microtubules from a defined point is essential. In branches of phylogeny that do not contain centrioles, cells use alternative strategies to organize their mitotic microtubules. Fungi have developed a structure that is functionally analogous to the centriole called the spindle pole body (Kubai,

1 Portions of this chapter were previously published in Cilia, 2016 Jan 19;5.1 PMID:26793300, and are included with the permission of the copyright holder.

1975). Additionally, higher land plants use their nuclear envelope to organize mitotic microtubules until nuclear breakdown, after which microtubules organize around kinetochore fibers (Palevitz, 1993). In branches of phylogeny that do not contain basal bodies, cilia and flagella do not form. CBBs are highly conserved phylogenetically and remain essential for microtubule organization in a majority of eukaryotic cells.

Centrioles recruit pericentriolar material (PCM) to make up centrosomes, the major eukaryotic microtubule organizing center. During mitosis the centrosome nucleates both astral and spindle microtubules. Astral microtubules are important during mitosis, where they attach to the cell cortex and position the spindle, essentially defining the geometry of the division plane. As such, loss of centriole components has detrimental effects on spindle orientation and profound effects on the fidelity of asymmetric cell divisions (Hyman, 1989;

Khodjakov and Rieder, 2001; Toyoshima et al., 2007). Spindle microtubules attach to chromosomes and use their dynamicity to pull chromosomes apart during metaphase. The effect of centriole loss on the ability to form bipolar spindles is complex. Early studies sought to understand the importance of centrioles by laser ablation (Khodjakov et al., 2000), microsurgical removal (Hinchcliffe et al., 2001; Maniotis and Schliwa, 1991), or antibody injection against centriolar components (Bobinnec et al., 1998a). The conclusion from these studies is that bipolar mitotic spindles can form in the absence of centrioles. Unfortunately, the methods used in these studies were only able to remove the centrioles and were unable to completely remove residual PCM so the effect of complete centrosome loss was not directly tested. To directly test this, further studies used genetic approaches to inhibit centriole duplication, which results in lack of PCM recruitment in subsequent cell cycles.

These cells are still able to form bipolar mitotic spindles, however, there is pronounced mitotic delay, chromosome instability and aneuploidy (Basto et al., 2006; Bettencourt-Dias et al., 2005; Sir et al., 2013). These results suggest that centrioles are essential for proper progression through mitosis and the fidelity of chromosome segregation.

Outside of mitosis centrioles function as basal bodies, which anchor and nucleate the microtubules of the flagellar and ciliary axoneme. Flagella are typically used in locomotion of prokaryotic and eukaryotic cells, although there is evidence that flagella act as sensory organelles as well (Silflow and Lefebvre, 2001; Wang et al., 2005). Cilia are multifunctional cellular appendages that extend into extracellular space. Depending on the cell type, cilia come in two main varieties, primary and motile. A single primary cilium is found in nearly every human cell and functions as a signaling center. Primary cilia play an integral role in developmental signaling pathways, most notably hedgehog signaling (Goetz and Anderson,

2010). Additionally, primary cilia can function as mechanosensors that transduce signal based on the bending of their axoneme caused by environmental fluid flow (AbouAlaiwi et al., 2009; Nauli et al., 2011; Nguyen and Jacobs, 2013; Roth et al., 1988). Motile cilia bend their axonemes as well, but unlike primary cilia, motile cilia bending is intrinsically controlled.

These epithelial cell cilia are typically found in arrays that undulate in a coordinated manner to move fluid in a single defined direction (Chilvers and O'Callaghan, 2000). Motile cilia are required for a number of essential processes in humans including movement of cerebral spinal fluid, clearance of mucus from the respiratory tract and progression of the egg down fallopian tubes (Lyons et al., 2006; Sawamoto et al., 2006; Wanner et al., 1996). Loss of either primary or motile ciliary function results in a diverse collection of developmental disorders termed ciliopathies (Hildebrandt et al., 2011). A number of ciliopathy genes encode basal body proteins, however, the majority of ciliopathy genes encode proteins that function at the cilium (Veleri et al., 2014; Williams et al., 2011). The enrichment of cilia- specific ciliopathy genes over basal body-specific ciliopathy genes can in part be explained by the fact that mutations that cause basal body defects are often lethal. Loss of basal bodies results in a complete loss of cilia, resulting in lethality in humans. Furthermore, basal body components are also centriolar components so defects in basal bodies that also affect centriole function may manifest as mitotic defects and not ciliopathies. Overall, basal bodies

are required for nucleation of flagella and both primary and motile cilia and loss of basal bodies results in severe disorders called ciliopathies.

Centriole and basal body structure

The foundation of our knowledge of CBB structure has been laid down by early electron microscopy studies of the 1960’s and 1970’s (Allen, 1969; Cavalier-Smith, 1974;

Dippell, 1967; Dippell, 1968; Dirksen, 1971; Sorokin, 1968a). These studies have shown that CBBs have remarkable structural conservation across phylogeny. The CBB is a cylindrical shaped structure that is composed of nine sets of modified triplet microtubules (A-

B-C-tubules) that are radially arranged around a cartwheel-like structure at its proximal end

(Figure 1.1). Over the proximal third of CBBs there are linkages between adjacent triplet microtubules called A-C linkers (Figure 1.1)(Allen, 1969). As the name suggests these linkages extend from the C-tubule of one triplet microtubule to the adjacent triplet microtubule A-tubule. Recently, advances in cryo-tomography has made it possible to study

CBBs in more detail than was previously available. Notably, these studies have identified new conserved structural domains within the triplet microtubules, which may be indicative of specific microtubule associated protein (MAP) binding (Li et al., 2012). Although highly conserved, there is some structural diversity that is apparent in the core CBB structure.

Every CBB has a cartwheel, and some species (including humans) lose their cartwheel after the initial assembly events (Vorobjev and Chentsov, 1980). Additionally, there are examples of CBBs that have varying numbers of microtubules instead of the canonical triplet structure.

Most notably, Drosophila have CBBs that contain doublet microtubules, and C. elegans contain CBBs with one microtubule (Gogendeau and Basto, 2010; Pelletier et al., 2006).

Another area of diversity seen in the CBB is its overall length. Typically the CBB ranges from

400-500 nm in length but some species have exceptionally long CBBs like in Drosophila

spermatocytes and Trichonympha, which harbor CBBs that are up to 1.3m and 5m long, respectively (Guichard and Gonczy, 2016; Tokuyasu, 1975).

Outside of its core structure, the CBB also contains structural domains that are required specifically for its function as either a centriole or a basal body. Centrioles recruit a structurally amorphous protein meshwork called the PCM to aid in the nucleation of microtubules (Nigg and Stearns, 2011).Though recent advanced microscopy techniques have revealed that the PCM is much more structured than previously thought, its overall structure remains elusive (Mennella et al., 2014). The PCM is only present in CBBs that have centriolar functions. In these cells the PCM remains associated at the centriole while the centriole functions as a basal body (Moser et al., 2010). In cells without centrioles, no

PCM is ever recruited to basal bodies. CBBs also contain structures required for basal body specific functions. At the distal end of CBBs is a transition zone, which is involved in creation of the ciliary vesicle and is required for ciliogenesis (Figure 1.1). Additionally, the distal end of CBBs contains two sets of cone shaped structures called the distal and subdistal appendages, which are required for docking the CBB at the plasma membrane. These basal body specific structures are present even when the CBB functions as a centriole. Emerging

evidence suggests that the subdistal appendages may have a secondary role in mitotic spindle positioning (Chen et al., 2014; Hung et al., 2016). The CBB has strong structural conservation, which is mostly maintained when altering between roles as a centriole and a basal body.

Molecularly, the CBB has a high amount of conservation across phylogeny

(Carvalho-Santos et al., 2010). The triplet microtubules that comprise the length of CBBs are primarily made up of - and -tubulin, and there are well established roles for -tubulin in assembly of new CBBs. Additionally - and -tubulin are implicated in the formation and stabilization of CBB microtubules (Dupuis-Williams et al., 2002; Dutcher et al., 2002;

Garreau de Loubresse et al., 2001; Moudjou et al., 1996; Ross et al., 2013; Shang et al.,

2002). Interestingly, organisms that lack - and -tubulin also lack triplet microtubules as is seen in subsets of Drosophila CBBs and C.elegans (Carvalho-Santos et al., 2010). This suggests that - and -tubulin are at least in part required for triplet microtubule formation.

The cartwheel is composed of of the protein Sas6 (Kitagawa et al., 2011). At the tips of each

“spoke” is the protein Bld10/Cep135, which binds both SAS6 and microtubules to connect the spoke to the A-tubule of a triplet microtubule (Hiraki et al., 2007). Bld10/Cep135 also bind the triplet microtubule associated protein CPAP/SAS4 (Lin et al., 2013). Together these three proteins, Sas6, Bld10/Cep135, and CPAP/SAS4 make up the ancestrally conserved

UNIMOD module (Carvalho-Santos et al., 2010). These three proteins are conserved in nearly every known species that contain CBBs and are critical for CBB assembly (see below) (Carvalho-Santos et al., 2010). Consistent with the idea that the cartwheel is an ancient structure, the cartwheel scaffolding protein STIL/SAS5 is among the most conserved

CBB proteins (Delattre et al., 2004). Outside of the cartwheel and the triplet microtubules, proteins are found to have high conservation but in a more taxon specific manner (Carvalho-

Santos et al., 2010). For example, most higher eukaryotes share a similar protein cascade

for initiating new CBB assembly (covered below). Also, CBBs that have functions as centrioles contain the PCM protein pericentrin (Doxsey et al., 1994). Overall, the molecular conservation of core proteins is remarkable and suggest a very ancient common origin for

CBBs.

CBBs are highly conserved both structurally and molecularly. With the diversity of cellular contexts found in eukaryotes it is striking that the CBB shows such high levels of structural and molecular conservation. The commonality between structure and function would suggest that CBBs have a common way in which they are assembled. UNIMOD proteins are essential for CBB assembly and are found in nearly every organism that assembles CBBs.

Centriole and basal body assembly

CBB assembly is tightly regulated. Centrioles duplication is tightly regulated.

Centrioles duplicate once and only once during the cell cycle, and like DNA this duplication occurs in S phase (Holland et al., 2010). The tight regulation of duplication is necessary because aberrant centriole number promotes genomic instability (Ganem et al., 2009;

Silkworth et al., 2009). Centriole biogenesis occurs off of an existing centriole in what is called the templated duplication pathway. During templated duplication, centrioles form off of the proximal end of an existing centriole though the exact mechanism of new centriole site determination is unknown. Templated duplication is not the only way to generate new CBBs.

In cells that form multi-ciliary arrays of motile cilia or in cells that have had their centrosomes ablated, basal bodies are generated through a different pathway called de novo pathway

(Anderson and Brenner, 1971; Dirksen, 1971; Kalnins and Porter, 1969; La Terra et al.,

2005; Sorokin, 1968a). Naturally occurring de novo assembly of basal bodies occurs in

G0/G1 phase of the cell cycle and occurs off of a matrix of fibrous granules called the deuterosome (Chang et al., 1979). Unlike templated duplication, a single deuterosome nucleates an average of 5.6 basal bodies at a time, thereby greatly increasing the amount

basal bodies being produced at a given time (Al Jord et al., 2014). Regulation of this pathway is less understood that that of the templated assembly pathway, however, recent research is shedding light on the initiation of this process (Klos Dehring et al., 2013; Zhao et al., 2013). Regardless of whether new CBBs are templated from existing CBBs or are generated de novo the CBB assembly pathway can be broken down into three phases: initiation, formation and elongation of triplet microtubules, and maturation.

Initiation of centriole duplication is a hierarchical process that involves coordination between a number of proteins. In the templated centriolar duplication pathway procentriole formation begins at the proximal end of the templating centriole in a position perpendicular to its length. First, SPD2/Cep192 and Asterless/Cep152 are recruited in tandem to the site of procentriole formation (Hatch et al., 2010; Kim et al., 2013; Sonnen et al., 2013). Together these proteins act as a scaffold for the recruitment of Plk4/SAK/ZYG-1 (Kim et al., 2013).

Plk4/SAK/ZYG-1 is a kinase that is largely considered to be the master regulator of centriole biogenesis due to the fact that it is necessary and sufficient for centriole duplication

(Bettencourt-Dias et al., 2005; Kleylein-Sohn et al., 2007). How exactly Plk4/SAK/ZYG-1 promotes centriole biogenesis is just now being identified. There is mounting evidence that

Plk4 dependent phosphorylation of STIL/SAS5 causes STIL/SAS5 to directly recruit and complex with UNIMOD protein Sas6 at the site of new assembly, now called the procentriole

(Arquint et al., 2015; Kratz et al., 2015; Lopes et al., 2015; Moyer et al., 2015). At this point

Sas6 dimerizes to create the canonical 9-fold cartwheel structure (Kitagawa et al., 2011).

Although Sas6 is found in every organism that contains a CBB, a clear homolog of

Plk4/SAK/ZYG-1 has not been identified in all systems that have templated centriole biogenesis suggesting that centriole duplication may be initiated differentially (Carvalho-

Santos et al., 2010).

After establishment of the cartwheel by Sas6, new CBBs are able to begin building the triplet microtubules. The triplet microtubules are added sequentially beginning with the

A-tubule and followed by the B- and C-tubules. Formation and attachment of the triplet microtubules to the cartwheel spokes requires both other UNIMOD proteins CPAP/SAS4 and Bld10/Cep135. CPAP/SAS4 is a microtubule binding protein that is stabilized by - tubulin during microtubule formation (Dammermann et al., 2008; Hsu et al., 2008).

CPAP/SAS4 is required for triplet microtubule formation and elongation (Schmidt et al.,

2009; Tang et al., 2009). Furthermore, overexpression of CPAP is sufficient to cause overly long CBBs (Kohlmaier et al., 2009). Bld10/Cep135 is not required for triplet formation but is required for attachment of the triplet microtubules to the cartwheel spokes (Hiraki et al.,

2007). Loss of Bld10/Cep135 results in structurally unstable CBBs (Hiraki et al., 2007). This finding suggests that during early assembly the cartwheel not only templates the 9-fold symmetry of CBBs but also scaffolds new assembly (Dahl et al., 2015; Hiraki et al., 2007;

Jerka-Dziadosz et al., 2010; Lin et al., 2013). Together all of the proteins required for CBB duplication initiation and elongation of triplet microtubules work together to create the core structure of the CBB.

After the triplet microtubules are assembled and elongated, the CBB can begin the process of maturation. Maturation is a process by which proteins are added to the completed CBB structure. These proteins are accessory to the core structure of the CBB and are required for the functional roles they serve. The most well understood examples of

CBB maturation are from mammalian centrioles. Centriole maturation is considered complete when a new centriole can act as a microtubule organizing center and can nucleate a primary cilium (Chretien et al., 1997). Because having multiple microtubule organizing centers or primary cilia is so detrimental to genome stability the recruitment of structural proteins is tightly regulated in a cell cycle dependent manner. Normal centriole maturation is a slow process, it takes upwards of two cell cycles to complete (Kong et al., 2014). To facilitate this maturation, centrioles must recruit PCM to gain microtubule organizing center functionality and form the distal and subdistal appendages required in plasma membrane

docking for cilium nucleation. Formation of these structures requires the accumulation of numerous proteins (Doxsey et al., 1994; Graser et al., 2007; Gromley et al., 2003;

Guarguaglini et al., 2005; Lange and Gull, 1995; Mogensen et al., 2000; Nakagawa et al.,

2001). PLK1 is considered a master regulator of centriole maturation because unregulated

PLK1 activity results in acceleration of centriole maturation as shown by accumulation of maturation markers (Kong et al., 2014). Furthermore, PLK1 has been shown to directly phosphorylate pericentrin, which in turn recruits a number of PCM components required for microtubule organizing center activity (Lee and Rhee, 2011). These findings support the idea that the CBB assembly is not complete until it is fully mature.

Relative to templated centriole assembly, much less in known about de novo basal body duplication. Recent studies have made clear that the same proteins (Cep192, Plk4,

Sas6) involved in templated centriole initiation are also necessary for de novo basal body duplication, suggesting that de novo basal body assembly may not be so different from what we already know about templated centriole duplication (Klos Dehring et al., 2013; Vladar and Stearns, 2007). Furthermore, there is no reason to believe that the proteins involved in elongation of the triplet microtubules are any different because de novo generated basal bodies still contain Bld10/Cep135 and CPAP/SAS4. Besides the nucleating structure, the major difference between templated centriole assembly and de novo assembly is that templated centriole duplication required more than a cell cycle to mature whereas basal bodies generated through de novo assembly can mature without undergoing cell division.

Maturation occurs quickly as newly formed basal bodies are able to nucleate a cilium without going through a cell cycle. This process includes the formation of distal and subdistal appendages. Overall, the presence of a non-templated pathway for basal body biogenesis suggests that basal bodies are able to self-assemble in the presence of differential initiation cues.

CBBs are large macromolecular machines. Over or under duplication of CBBs is detrimental to the cell. Because CBBs have self-assembling capabilities its assembly must be highly organized and regulated. The process of CBB assembly is complex, and although originally thought to be hierarchical, and the proteins involved in the progression of assembly are interdependent on each other. Although two different pathways exist to create

CBBs there appears to be significant overlap between the two. Furthermore, loss of early assembly proteins results in structural instability suggesting that proper assembly is critical for CBB function (Jerka-Dziadosz et al., 2010).

Conclusions

CBBs are ancient macromolecular structures that act as microtubule organizing centers in eukaryotic cells. As centrioles they nucleate the mitotic spindle and as basal bodies they nucleate the microtubules of the ciliary axoneme. Their function is essential for organism viability. In order to function they utilize a catalog of conserved proteins that impart the canonical 9-fold symmetry of their core structure. These proteins include a conserved module (Sas6, Bld10/Cep135, and CPAP/SAS4) termed UNIMOD, which are essential for assembly of the cartwheel and triplet microtubules. Initiation of CBB assembly is a tightly regulated process that involved the cooperation of many different proteins. Loss of early assembly proteins compromises the structural integrity CBBs suggesting that proper assembly of CBBs is required for its structural integrity.

Centriole and Basal Body Stabilization

CBBs are structures that experience tremendous amounts of mechanical force. The centrosome organizes the microtubules necessary for mitosis. These microtubules emanate from -tubulin seeds embedded in the centrosomal PCM. It is clear that the pulling forces generated by spindle microtubules are transferred to the centriole because centriole integrity is necessary to prevent centrosome fragmentation during mitosis (Abal et al., 2005). In addition to spindle microtubules, astral microtubules also produce force on the centrosome.

During spindle pole orientation the centrosome is positioned by astral microtubule contacts with the cell cortex (Kozlowski et al., 2007). This positioning is mediated by cortical dyneins that are thought to function by holding disassembling microtubules in place (Nguyen-Ngoc et al., 2007). The force generated by a single disassembling microtubule is on the order of

~50pN, which is comparable to the pulling forces experienced on astral microtubules during centrosome positioning (Grill et al., 2003; Grishchuk et al., 2005). Failure to stabilize centrioles from mechanical forces causes centrosome fragmentation, which leads to aneuploidy, a hallmark of cancer (Bastians, 2015). Basal bodies experience different types of mechanical forces. They experience compression, tensile, and shear forces generated from the movement of the cilia and flagella that they nucleate (Bayless et al., 2012; Galati et al., 2014; Lindemann and Kanous, 1997; Riedel-Kruse et al., 2007; Vernon and Woolley,

2004). This is especially apparent in basal bodies that nucleate and anchor motile cilia or flagella where the cilia actively beat. Although the types of forces experienced by centrioles and basal bodies are different, the structure is maintained by similar mechanisms. First,

CBBs use proteins to create essential linkages between structural domains. Second, CBB microtubules are stabilized by their triplet structure, post-translational modifications, and

MAPs.

Structural stabilization

CBBs need to be held together tightly if they are to resist the mechanical forces exerted on them. To ensure that their structural integrity remains intact, CBBs make connections between its cartwheel and the triplet microtubules, triplet microtubules to adjacent triplet microtubules, and their overall structure to their environment. Each of these connections is essential to stabilize the structure from the forces it routinely experiences.

The CBB cartwheel not only organizes its nine-fold symmetry but is also necessary to stabilize the CBB from mechanical forces (Izquierdo et al., 2014). Because the cartwheel is lost from centrioles after assembly it has long been thought that the cartwheel is only

essential as a scaffold for early assembly of CBBs (Alvey, 1986; Gonczy, 2008; Vorobjev and Chentsov, 1980). However, recent work identified that centrioles must have either a cartwheel or PCM to remain intact through mitosis, suggesting that maturation to a centrosome is required before centrioles lose the cartwheel (Izquierdo et al., 2014).

Additionally, basal bodies of motile ciliary arrays and flagella, which do not generate PCM, do not lose their cartwheels, suggesting that the presence of a cartwheel may be essential to maintain the structure in the face of ciliary/flagellar beating forces (Allen, 1969; Dippell,

1967; Vladar and Stearns, 2007). The cartwheel typically resides at the proximal end of

CBBs and it can vary in size, which may affect its ability to stabilize CBBs. In fact, the size of the cartwheel can change in length throughout the cell cycle. In the case of Trichonympha, a single celled flagellate, the cartwheel can extend ~50 times longer up the length of the basal body than that of centriolar cartwheels (Guichard et al., 2012; Guichard and Gonczy, 2016).

Since Trichonympha live in an extremely viscous host environment, the additional cartwheel length may be necessary to resist additional mechanical forces associated with moving a flagella through a viscous environment (Jung et al., 2014; Spoon et al., 1977). Overall, the cartwheel is necessary for assembly of CBB and its presence may help to maintain CBB structural integrity.

In order for the cartwheel to stabilize CBBs their spokes must make proper attachments to the triplet microtubules. This attachment is mediated by the UNIMOD protein

Bld10/Cep135 (Hiraki et al., 2007; Matsuura et al., 2004). Bld10 mutants result in loss of triplet microtubules and ultimately entire basal bodies in the model systems Paramecium,

Chlamydomonas, and Tetrahymena (Bayless et al., 2012; Hiraki et al., 2007; Jerka-

Dziadosz et al., 2010). Furthermore, the loss of basal bodies in Tetrahymena Bld10 mutants is completely rescued by inhibition of ciliary beating (See chapter II) (Bayless et al., 2012).

This finding suggests that mechanical forces exerted on the basal body by cilia are sufficient

to cause structural disassembly when the basal body consists of weakened structural attachments.

The CBB also has structural attachments between neighboring triplet microtubules.

These attachments appear in the proximal third of CBBs as electron dense fibers that extend from the A-tubule of one triplet microtubule to the C-tubule of an adjacent triplet microtubule and are termed A-C linkers (Gibbons and Grimstone, 1960; Guichard et al.,

2013; Li et al., 2012). The composition of A-C linkers is not well known, however recent work has shown that loss of the highly conserved ciliopathy protein Poc1 results in severely shortened or lost A-C linkers (Meehl, 2016). Furthermore, this loss renders basal bodies susceptible to cilia generated forces resulting in loss of triplet microtubules and ultimately entire basal bodies (Meehl, 2016; Pearson et al., 2009b). Importantly, Poc1 also plays a role in centriole stabilization, which indicates that the same structural connections necessary for basal body stabilization may be used in centriole stabilization (Venoux et al., 2013). Overall, these results suggest that, like the attachments between the cartwheel and the triplet microtubules, attachments between adjacent triplet microtubules are necessary to stabilize

CBBs from mechanical forces.

In addition to connections within itself, CBBs create connections with their environment to help anchor and stabilize themselves. Centrioles form attachments to their

PCM through a hierarchical ordering of a number of proteins (Lawo et al., 2012). These attachments suggest that forces placed on centrosomes during mitosis are distributed across the entire structure, which may explain why PCM is so important for centriole integrity

(Izquierdo et al., 2014). Through astral microtubules, centrosomes form connections with the cell cortex that may further help stabilize them from pulling forces. Basal bodies do not have to resist forces during mitosis but they do need to resist forces derived from ciliary movement. To do this basal bodies utilize distal appendage proteins to connect to the plasma membrane. Docking the basal body at the plasma membrane is necessary for

ciliogenesis, but it is unknown whether distal appendage mediated attachments to the plasma membrane are necessary for basal body stabilization from ciliary movement. Basal bodies of motile cilia and flagella presumably experience additional mechanical forces compared with those that nucleate primary cilia. Interestingly, basal bodies are equipped with accessory structures that are necessary for their organization and stability. Motile cilia and flagellar basal bodies have unique accessory structures that form connections to their environment (Pearson, 2014). The most phylogenetically conserved of these accessory structures is the striated fiber. As its name implies, this structure is an electron dense striated fiber that connects basal bodies to the plasma membrane (Allen, 1969; Holmes and

Dutcher, 1989; Iftode and Fleury-Aubusson, 2003; Jerka-Dziadosz et al., 1995; Lechtreck and Melkonian, 1991; Sperling et al., 1991; Zhang and Mitchell, 2015). Striated fibers are made up in part by SF-Assemblin, a coiled-coil protein that is able to form striations in vitro

(Lechtreck and Melkonian, 1998; Weber et al., 1993). Importantly, truncations of the striated fibers lead to basal body orientation and stability defects, suggesting that connections between the basal body and its environment may act to displace the mechanical forces it experiences (Galati et al., 2014).

CBBs use structural components to form the connections between their own domains and their environment to maintain stability from the mechanical forces that they experience. The cartwheel itself acts as a CBB stability factor (Izquierdo et al., 2014). In order to stabilize CBBs the cartwheel makes spoke to triplet microtubule connections mediated by Bld10/Cep135 (Bayless et al., 2012; Hiraki et al., 2007; Jerka-Dziadosz et al.,

2010). Another way CBBs stabilize their structure is through triplet microtubule-triplet microtubule connections. These connections are mediated by A-C linkers that are, at least in part, composed of the conserved ciliopathy protein Poc1 (Figure 1.1)(Meehl, 2016). Outside of CBB domain linkages the CBB also uses connections with its environment to stabilize itself (Kozlowski et al., 2007; Pearson, 2014). Overall, structural connections and

environmental connections are indispensable for maintenance of basal body integrity in the face of mechanical forces.

Triplet microtubule stabilization

Microtubules are polymers that exhibit inherent and persistent growth and shrinkage

(Mitchison and Kirschner, 1984). The triplet microtubules of CBBs are unique microtubules in that they display no detectable dynamics outside of their initial assembly. Additionally, these microtubules are known to be extremely stable. The tubulins of the triplet microtubules exhibit minimal turnover dynamics (Kochanski and Borisy, 1990; Pearson et al., 2009a).

Triplet microtubules are resistant to typical microtubule stressors such as cold temperature, microtubule poisons and mechanical forces (Abal et al., 2005; Brinkley and Cartwright,

1975; Fracek and Margulis, 1979; Pearson et al., 2009b). In order to preserve CBB functionality, the triplet microtubules must regularly resist mechanical forces. CBB triplet microtubules resist these forces through their unique triplet structure, presence of microtubule associated proteins, and tubulin post-translational modifications.

The core structure of the CBB is composed of modified triplet microtubules that are necessary for their unique stable properties. These microtubules are composed of a complete 13 protofilament A-tubule followed by two successive 10 protofilament tubules (B- and C-tubules) that share a protofilament wall with it’s neighboring tubule (Figure 1.1)

(Guichard et al., 2013; Li et al., 2012). Ciliary axonemes consist of doublet microtubules, which must be flexible, yet stable enough, to bend without breaking. It is possible that CBB triplet microtubules use the additional microtubule to increase stability. The presence of triplet microtubules is strongly correlated with whether an organism encodes the  and  isoforms of tubulin, suggesting that differential tubulin isoforms may be involved in formation of tubulin triplets (Carvalho-Santos et al., 2011). -tubulin was first identified in

Chlamydomonas and when mutated in Chlamydomonas or knocked down in Paramecium it

leads to the loss of the majority of basal body C-tubules (Dutcher and Trabuco, 1998;

Garreau de Loubresse et al., 2001; O'Toole et al., 2003). Ultimately, loss of C-tubules caused by -tubulin knock down in Paramecium results in unstable basal bodies that fall apart, suggesting that they are innately unstable or not able to withstand the forces experienced by ciliary beating (Garreau de Loubresse et al., 2001). Genetic studies have found that -tubulin interacts with -tubulin, which may correspond to cryo-electron microscopy findings of a unique linker between the B- and C-tubules (Fromherz et al., 2004;

Li et al., 2012). -tubulin was first identified in humans and is essential for centriole duplication (Chang et al., 2003; Chang and Stearns, 2000). Subsequent studies in

Chlamydomonas and Paramecium also find -tubulin to be essential for centriole assembly

(Dupuis-Williams et al., 2002; Dutcher et al., 2002). Notably, in Tetrhaymena, -tubulin loss in existing basal bodies causes a progressive loss of B- and C-tubules culminating with loss of entire basal bodies (Ross et al., 2013). Non-tubulin basal body stability factors also show loss of B- or C-tubules, most notably the junction domain linking proteins Bld10 and Poc1

(Bayless et al., 2012; Meehl, 2016). These findings suggest that maintenance of triplet microtubule structure is essential for CBBs ability to resist mechanical forces.

The triplet microtubules of CBBs are decorated with post-translational modifications and these modifications are necessary for CBB stability. Tubulins can be post-translationally modified in many ways (acetylation, tyrosination, glutamylation, glycylation), and each of these modifications affects the dynamics of microtubule growth and stability (Wloga and

Gaertig, 2010). Tubulin acetylation is thought of as a hallmark of stable populations of microtubules as it is found to be enriched in neurons, cilia, and CBBs (Black and Keyser,

1987; Kim, 1991; Loktev et al., 2008; Maruta et al., 1986; Matsuyama et al., 2002). Of the common post-translational modifications of tubulin, acetylation is the only one whose major modification site (K40 -Tubulin) resides in the lumen of the microtubule (Nogales et al.,

1998). Although no mechanism for how acetylation of -tubulin affects microtubule stability has been determined, recently it has been hypothesized that acetylation could increase tubulin protofilament spacing, which in turn could allow greater microtubule plasticity (Howes et al., 2014). This plasticity is desired in microtubules that experience bending and compressive forces such as those found in cilia and CBBs.

Most other tubulin post-translational modifications are found on the carboxy-terminal tails of both - and -tubulin (Wloga and Gaertig, 2010). Both - and -tubulin carboxy- terminal tails are highly conserved regions of less than 40 amino acids that are dynamic at the exterior of the microtubule (Chakraborti et al., 2016). Post-translational modifications at carboxy-terminal tails may promote or repel MAP binding. Tubulins of the ciliary axoneme and CBBs are enriched for carboxy terminal tail associated post-translational modifications glutamylation and glycylation (Bobinnec et al., 1998b; Bre et al., 1994; Bre et al., 1996; Wolff et al., 1992). These modifications compete to form chains off of the same glycine residues

(Wloga and Gaertig, 2010). The addition of glycine or glutamate residues is catalyzed by the tubulin tyrosine ligase-like (TTLL) family of enzymes (Janke et al., 2005; Wloga et al., 2009).

Overexpression, knockdown and knockout studies have assessed these enzymes with respect to the function of glycylation and glutamylation. In ciliary axonemes tubulin glycylation promotes microtubule stability (Bosch Grau et al., 2013; Pathak et al., 2011;

Wloga et al., 2009). Tubulin glycylation function has not yet been assessed with regard to

CBBs, though it is an intriguing avenue for future research. Functional studies on tubulin glutamylation reveal a more complex function. Overexpression of TTLL6, a tubulin glutamylase, shows that tubulin glutamylation can both stabilize and destabilize microtubules within the same cell (Wloga et al., 2010). In these cells axnonemal microtubules are destabilized and cytoplasmic microtubules are stabilized by glutamylation.

Axoneme destabilization is facilitated by katanin mediated severing of microtubules, which is

targeted to glutamylated doublet microtubules (Lacroix et al., 2010; Sharma et al., 2007).

Cytoplasmic microtubule stabilization is mediated by an unknown mechanism. Interestingly,

CBBs are specifically glutamylated by two TTLL family members, TTLL1 and TLL9 (Wloga et al., 2008). Specific loss of basal body glutamylation, through TTLL1 and 9 knockout, results in destabilized basal bodies, suggesting that basal body glutamylation is a stabilizing modification (see Chapter III). The differential effects of tubulin glutamylation may be indicative of different MAPs that rely on glutamylation for targeting of different microtubule populations. Intriguingly, antibodies that bind glutamylated tubulin cause centriolar disassembly in HeLa cells, which may be indicative of blockage of MAP binding (Bobinnec et al., 1998a). Overall, tubulin post-translational modifications are important for maintaining the triplet microtubules through unknown mechanisms that may rely on MAP binding.

The CBB triplet microtubules are bound by a number of MAPs. Proteins that bind the triplet microtubules are typically found to be microtubule stabilizing proteins. CAP350 is a microtubule binding protein that interacts with the microtubule plus end binding protein EB1 at centrioles to aid in centrosomal microtubule anchoring (Yan et al., 2006). In addition to anchoring centrosomal microtubules CAP350 directly stabilizes the centriolar triplet microtubules (Le Clech, 2008). Consistent with a role in microtubule stabilization CAP350 stabilizes microtubules associated with the Golgi apparatus (Hoppeler-Lebel et al., 2007).

Fop1 interacts with CAP350 at centrioles (Yan et al., 2006). In Tetrahymena, Fop1 associates along the length of the triplet microtubules and is necessary for basal body stabilization against ciliary beating generated mechanical forces (see Chapter III). The CBB structural domain connecting proteins Bld10 and Poc1 both bind microtubules and have microtubule stabilizing capabilities (Carvalho-Santos et al., 2012; Venoux et al., 2013).

Together, these findings suggest that CBBs are stabilized by microtubule binding proteins and their targeting to the triplet microtubules may be influenced by tubulin post-translational modifications.

Conclusions

CBBs are structures that need to resist mechanical forces to function. To resist these forces CBBs are stabilized in a number of ways. Their overall structure is held together by domain connections. The cartwheel itself is a stabilizing structure and the Bld10/Cep135 mediated connections of the cartwheel spokes to the triplet microtubules are necessary to resist mechanical forces (Bayless et al., 2012; Izquierdo et al., 2014). Triplet microtubules are also connected to each other through A-tubule to C-tubule linkers. Loss of these linkers renders CBBs susceptible to mechanical forces (Meehl, 2016). CBBs are kept anchored by connections to their environment. For centrioles, this can be mediated by PCM and for basal bodies these connections rely on a number of accessory structures, notably the striated fiber

(Izquierdo et al., 2014; Kozlowski et al., 2007; Pearson, 2014). CBBs also take measures to ensure that their triplet microtubules are stabilized. The CBB triplet microtubules that extend to form the length of the CBB are uniquely stable. Their triplet structure imparts stability and is reliant on and tubulin (Carvalho-Santos et al., 2011). Outside of their triplet structure,

CBB microtubules are stabilized by post-tranlsational modifications (acetylation, glycylation and glutamylation) and number of microtubule associated proteins (Bayless et al., 2012;

Carvalho-Santos et al., 2012; Le Clech, 2008; Venoux et al., 2013; Wloga and Gaertig,

2010). In summary, CBBs utilize their structure and their environment to resist mechanical forces.

Motile Cilia and Flagellar Beating

Centrioles and basal bodies share most of the same structural components. Loss of function mutations of these components often results in mitotic defects and cell cycle arrest due to the centriole’s major role in facilitating the progression of mitosis (Cheng et al., 2008;

Mikule et al., 2007). Additionally, many cell cycle checkpoint regulators localize to centrosomes, so disruption of centriole structural integrity can cause mitotic arrest (Jackman et al., 2003; Matsumoto and Maller, 2004). The prevalence of mitotic defects during CBB

mutagenesis can mask the finer function of many CBB components. This is especially true of CBB stability factor mutants that cause deterioration of CBBs over time as unmitigated mechanical forces result in accumulation of structural defects. Motile cilia and flagella are nucleated by basal bodies that must anchor and resist the mechanical forces generated by their beating. Importantly, motile cilia and flagellar basal bodies do not act as centrioles.

Because of this, and the direct measurable force produced by motile cilia or flagellar beating, the basal bodies of motile cilia and flagella serve as a great system to understand how CBBs resist mechanical forces.

Axoneme structure

The axoneme of motile cilia and flagella is primarily made up of tubulin. Axonemes are composed of nine sets of radially arranged doublet microtubules that are continuous with the A- and B- tubules of the basal body (Figure 1.2)(Afzelius, 1959; Fisch and Dupuis-

Williams, 2011). Thus any forces exerted on the axoneme are transmitted to the basal body.

In the center of the axoneme reside two microtubules called the central pair (Figure 1.2).

These two microtubules are not found in primary or nodal cilia but instead are only found in motile cilia and flagella (Fisch and Dupuis-Williams, 2011). Unlike CBBs, ciliary and flagellar axonemes only contain the canonical - and -tubulins (Mohri et al., 2012). However, there is evidence that a single axoneme utilizes multiple isoforms of - and -tubulin (Kobayashi and Mohri, 1977; Sullivan, 1988). It is thought that the axoneme utilizes these different isoforms for the formation of the B-tubules of the axoneme (Sullivan, 1988). Like CBBs, ciliary and flagellar axonemes are heavily modified by post-translational modifications including acetylation, glutamylation and glycylation (Huitorel et al., 2002; Wloga and Gaertig,

2010). Both glutamylation and glycylation of tubulin play a significant role in the assembly and maintenance of axoneme length as mutations result in shorter axonemes (Ikegami et al., 2010; Janke et al., 2005; Pathak et al., 2011; Wloga et al., 2009). Tubulin glutamylation

is also necessary for modulating the waveform of axonemal bending during flagellar beating

(Kubo et al., 2012; Kubo et al., 2010). Overall, the axonemal structure of motile cilia and flagella is composed of - and -tubulin arranged into nine radially symmetric doublets that connect with the triplet microtubules of their basal body. These tubulins are post- translationally modified and, like CBBs, these modifications are necessary for structural integrity of the axoneme.

Besides doublet microtubules, another major structural component of ciliary/flagellar axonemes are two protrusions that form off of the A-tubules and extend towards the B- tubule of adjacent microtubule doublets (Figure 1.2)(Afzelius, 1959). These outer and inner

“arms” off of the doublet microtubules are made up of the ATPase dynein, first identified in

Tetrahymena cilia extracts (Gibbons, 1963; Gibbons and Rowe, 1965; Ogawa et al., 1977;

Ogawa and Mori, 1975). Outer dynein arms are necessary for motile cilia and flagellar beating (see below). Outer dynein arms are thought to consist of a single type of dynein arranged in 24 nm intervals along the length of the axoneme (Inaba, 2007). In metazoans, this dynein consists of two heavy chains, three to five intermediate chains and six light chains (Gibbons, 1981). In Chlamydomonas, where a large majority of our knowledge about axonemal dynein comes from, the outer dynein arms consist of three heavy chains, two intermediate chains, and ten light chains (DiBella and King, 2001). The heavy chains of

outer dynein arms contain its ATPase activity and are necessary for microtubule sliding in both cilia and flagella (Moss et al., 1992; Tang et al., 1982; Toyoshima, 1987a; Toyoshima,

1987b; Yano-Toyoshima, 1985). The intermediate chains of the outer dynein arms are necessary for dynein assembly and binding to the A-tubule (Ogawa et al., 1996). The light chains of outer dynein arms are thought to be required for assembly and regulation of dynein motor function (Hozumi et al., 2006). Unlike outer dynein arms, inner dynein arms consist of multiple types of dyneins, up to seven in Chlamydomonas (Kagami et al., 1990).

Much less is known about inner dynein arm function but they are thought to be required for the waveform of the ciliary and flagellar beat stroke (Brokaw and Kamiya, 1987). Overall, outer and inner dynein arms are major structural components of axonemes and are required for the beating and waveform of motile cilia and flagella.

The last major component of ciliary/flagellar axonemes are radial spokes. Radial spokes are large multiprotein complexes made up of as many as 23 proteins in the case of

Chlamydomonas (Yang et al., 2006). Radial spokes are localized inside the doublet microtubules and have functions in regulating dynein activity (Figure 1.2). They are not thought to be necessary for axonemal bending and bend propagation but they are involved in controlling the properties of bend and modulating the overall beating frequency (Smith and Yang, 2004; Yang et al., 2006). Radial spokes are able to regulate dynein by attaching to inner dynein arms. Radial spoke proteins may stimulate dynein activity through

Ca2+/CaM-dependent or cAMP-dependent signaling as a number of its proteins function in these signaling pathways (Yang et al., 2006). In Chlamydomonas, radial spokes are attached to inner dynein arms through a multi-protein intermediate called the dynein regulatory complex, though no such structural homolog has been identified in metazoans

(Gardner et al., 1994; Heuser et al., 2009; Piperno et al., 1992). This complex contains proteins that form inter-doublet links that are thought to contribute to the elastic resistance necessary for converting microtubule sliding to axonemal bending (Heuser et al., 2009).

Ultimately, radial spoke proteins represent a third major component of the ciliary/flagellar axoneme that, along with accessory proteins, connect to inner dynein arms and aid in the creation of the waveform of axonemal bending.

Motile cilia and flagella are composed of an axoneme that is built to generate mechanical force. The axoneme is made up of nine doublet microtubules that are structurally attached to the basal body A- and B-tubules (Afzelius, 1959). Along the length of these doublets are two sets of dynein appendages. Outer dynein arms regulate the bending of the axoneme by attaching to neighboring doublet microtubules and walking along their length to create inter-doublet sliding (Moss et al., 1992; Tang et al., 1982; Toyoshima,

1987a; Toyoshima, 1987b; Yano-Toyoshima, 1985). Inner dynein arms regulate the waveform properties of axonemal bending by interaction with radial spoke proteins (Brokaw and Kamiya, 1987; Smith and Yang, 2004; Yang et al., 2006). Overall, the structural components of the motile cilia/flagellar axoneme are essential for propagation of axonemal beating.

Beating mechanism

Motile cilia and flagella beat by the coordinated bending of their axonemes. Original hypotheses for how axonemes bend proposed that the doublet microtubules shrink and grow relative to each other to produce curvature. The origin of axoneme curvature is determined by axonemal bending that is facilitated by inter-doublet microtubule sliding

(Satir, 1965; Satir, 1968). ATP is sufficient to cause doublet sliding in sea urchin sperm

(Summers and Gibbons, 1971). Using ATP to activate microtubule sliding in Tetrahymena axonemes shows that activated outer dynein arms on A-tubules of a doublet microtubules push the adjacent B-tubule towards the ciliary tip (Sale and Satir, 1977). This study was repeated with sea urchins and mammalian sperm flagella and the sliding went in the opposite direction (Ishijima et al., 1996; Lorch et al., 2008). This finding suggests that inter- doublet microtubule sliding can be altered based on the cellular context.

The sliding of doublet microtubules is influenced by a number of factors. These factors include the number of dynein arms on the doublet microtubule, and concentrations of

ATP, Ca2+ and cAMP. The velocity of microtubule sliding is proportional to the number of dynein arms present (Hata et al., 1980; Yano and Miki-Noumura, 1980). Sliding velocity is also directly proportional to the amount of ATP in the system. Oversaturating the system with ATP results in a maximum sliding velocity of 14 m/sec (Takahashi et al., 1982; Wada et al., 1991). Ca2+ causes a decrease in sliding velocity (Bannai et al., 2000; Nakano et al.,

2003). Interestingly, Ca2+ concentration influences the pattern of doublet microtubule sliding.

Low concentrations of Ca2+ cause doublet microtubule to act independently of one another and slide separately; however, high concentrations of Ca2+ result in doublet microtubules bundling together to create two groups of doublet microtubules that only slide relative to the other group (Ishijima et al., 1996; Nakano et al., 2003; Sale, 1986). The latter situation is what is seen in nature and results in the two sided beat of motile cilia and flagella. cAMP is not required for microtubule sliding and has no effect on the velocity of microtubule sliding, but it is required for flagellar bending in mammalian sperm (Kinukawa et al., 2006). The separation between microtubule sliding and ciliary/flagellar bending has been studied more extensively and it was determined that microtubule sliding is converted to ciliary/flagellar bending when there is resistance to microtubule sliding (Fujimura and Okuno, 2006;

Shingyoji et al., 1977). These findings demonstrate the need for a fixed end or physical connections between adjacent doublets to generate ciliary/flagellar bending. In vivo, the fixed end of the axoneme resides at basal bodies. Overall, the bending of axonemes to produce ciliary/flagellar beating is produced by sliding between the axonemal doublet microtubules. Inter-doublet sliding is triggered by dynein activity and is controlled by a number of factors including, number of dynein arms, ATP, Ca2+ and cAMP concentration.

The regulation of microtubule sliding in cilia and flagella must be modulated to account for the variation between the ciliary and flagellar beat stroke. Flagella beat along a

two-dimensional plane with equal force generated in each direction. This beat stroke is maintained by the inter-doublet microtubule sliding between opposing bundles of doublet microtubules, which effectively split the axoneme into two halves. Each half is bundled together by dynein bridges (Lindemann et al., 1992). Computer simulations have been created to model the flagellar beat stroke; however, none of these models are widely accepted (Lindemann et al., 1992; Riedel-Kruse et al., 2007; Woolley, 2010). This could be due to the emerging data that the flagellar beat stroke has three-dimensional components to its beat path that do not agree with computer simulations demonstrating a completely two- dimensional beat path (Ishijima, 2012; Ishijima et al., 1992; Ishijima and Hamaguchi, 1993;

Woolley and Vernon, 2001). The differences between the visual two-dimensional beat path and the modeling data, which suggest a three-dimensional beat path imply that there may be additional regulation to alter the beat path (Ishijima, 2012). One possible regulator of planar beating is the central pair complex. Evidence for this includes the fact that nodal cilia beat exclusively in a helical pattern and they do not contain a central pair (Shishikura and

Sekiguchi, 1979). Intriguingly, the stiffness of sea urchin sperm is significantly greater perpendicular to the beating plane compared to parallel to the beating plane (Ishijima and

Hiramoto, 1994). Much less is known about the regulation of the ciliary beat stroke and by all accounts it is more complex than that of the flagella. Cilia beat along a three-dimensional path that can be broken down into two phases: a power stroke and a recovery stroke. The power stroke is a two-dimensional force generating movement of the cilium. After the power stroke concludes, a three-dimensional recovery stroke rotates the cilium back into position for another power stroke. Computer modeling has recreated the Paramecium ciliary beat stroke and this work shows that two opposing sets of four microtubule doublets affect sliding against each other during the power stroke and recovery stroke, respectively (Sugino and

Naitoh, 1982). The remaining set of doublet microtubules does not have any role in microtubule sliding (Sugino and Naitoh, 1982). The activation of a specific half of the

axoneme is likely a mechanism used in flagellar beating as well as ciliary beating. As is seen in flagella, the motile cilia central pair microtubule likely contribute additional stiffness along the plane perpendicular to the power stroke, however it is unknown how the recovery stroke is affected by the fixed position of the central pair of microtubules. In summary, the beat stroke of flagella and motile cilia is complex and we are only now, through computer modeling and experimental imaging, beginning to understand the regulatory mechanisms that must be involved in coordinating these two different strokes.

Mechanical forces experienced at the basal body

The sliding of doublet microtubules is necessary to create the beat stroke of flagella and motile cilia. Doublet microtubules generate the force necessary to bend the axoneme by a stable attachment to the basal body while sliding. The result of this mechanism is that upward and downward forces are exerted on the basal body in the form of a shear force that can be broken down into compression and tensile forces (Riedel-Kruse et al., 2007; Vernon and Woolley, 2004). Additionally, ciliary/flagellar beating causes the basal body to rock back and forth. This movement causes the basal body to experience translational forces (Bayly et al., 2011). The differences between flagellar and motile ciliary beat strokes also suggest that basal bodies in these two systems experience mechanical forces differentially and subsequently may need to be stabilized differentially.

Compression and tensile forces are experienced when a system is compressed or stretched along a two-dimensional axis, respectively. These forces are predicted to be the greatest that the basal body experiences during ciliary/flagellar beating (Riedel-Kruse et al.,

2007). Basal body compression occurs when the doublet microtubules of the axoneme slide down relative to the plane of the transition zone (Vernon and Woolley, 2002; Vernon and

Woolley, 2004). In the context of an axonemal beat stroke, basal body compression occurs along the face of the basal body that is positioned towards where the power stroke terminates (Riedel-Kruse et al., 2007; Vernon and Woolley, 2004). The amplitude of

downward sliding and subsequent basal body compression has been visualized and measured in rat sperm to be 110 nm (Riedel-Kruse et al., 2007). This amount of compression represents about twenty percent of the total length of the basal body in this system. At the same time that one half of the basal body is experiencing compressive force the opposite face of the basal body is predicted to experience a stretching of tensile force that is equal in magnitude to the compressive force (Riedel-Kruse et al., 2007). Tensile forces at the basal body have not been measured or visualized yet and may be an intriguing area of future research. It is understood that the A- and B-tubules are continuous between the axoneme and the basal body, so compressive and tensile forces should translate directly between these regions. However, the exact amount of compressive and tensile force experienced by the basal body is difficult to model because the nature of the transition zone between the axoneme and the basal body is not well understood. There are a large number of proteins that localize to this region and some of them may act to dampen force in ways that we do not fully understand. Ultimately, compression and tensile forces represent the major forces experienced by basal bodies due to ciliary/flagellar beating.

In addition to compression and tensile forces, the basal body experiences translational forces from the back and forth rocking that occurs while the axoneme is moving

(Bayly et al., 2011). How these forces affect basal body stability is unclear, but the fact that the basal body has accessory structures that anchor and potentially reduce this rocking suggests that these forces must also be stabilized. This has been characterized in the greatest detail in Tetrahymena where the basal body accessory structures are well defined

(Allen, 1969). Tetrahymena kinetodesmal fibers connect the base of the basal body to the plasma membrane. When ciliary beating frequency is increased the kinetodesmal fiber extends and increases its contacts with the plasma membrane (Galati et al., 2014). A mutant that causes reduced kinetodesmal fiber length results in basal bodies that are unstable and oriented in obscure directions (Galati et al., 2014). These findings suggest that

translational forces are sufficient to move and destabilize the basal body in the absence of stabilizing structures.

Flagella and cilia have different beat strokes. The difference between these strokes has implications on how force is experienced at the basal body. Flagella beat along a largely two-dimensional plane. The force generated by the flagellar power stroke is equal in magnitude no matter what direction it is traveling (Riedel-Kruse et al., 2007). Thus, the flagellar basal body experiences an equal amount of compressive and tensile force along each face of the basal body. Motile cilia beat with the goal of moving fluid in a single direction. To achieve this they beat asymmetrically. The ciliary power stroke produces a large amount of force relative to the recovery stroke. As such, the microtubule sliding that occurs during the power stroke is predicted to be greater than during the recovery stroke.

This asymmetry in beating translates to a predicted asymmetry of forces experienced at the basal body. Compressive forces at the basal body would be greater along the side of the basal body that is facing the direction of power stroke movement. Conversely, tensile forces are greater along the side of the basal body that faces the direction of the recovery stroke.

In Tetrahymena, a system that displays a classical ciliary beat stroke, knockout of stability protein Poc1 causes disassembly of basal bodies along the side of the basal body that experiences greater compression forces (Meehl, 2016). This finding suggests that either compressive forces need to be stabilized to a greater degree than tensile forces or that

Poc1 specifically stabilizes the basal body against compressive forces. Overall, motile cilia are an intriguing model for the study of how CBBs resist mechanical forces because the different forces they experience can be separated and tracked along specific structural domains.

Conclusions

Ciliary and flagellar beating is a promising system to model how CBBs resist mechanical forces. The ciliary/flagellar axoneme consists of nine doublet microtubules

decorated with inner and outer dynein arms (Afzelius, 1959; Fisch and Dupuis-Williams,

2011). Sets of doublet microtubules are bundled together by dynein bridges to create two opposing sides of the axoneme (Lindemann et al., 1992). The activation of outer dynein arms causes each side to slide up and down relative to each other (Ogawa et al., 1977;

Ogawa and Mori, 1975). Activation of inner dynein arms translates that sliding into axonemal bending (Brokaw and Kamiya, 1987). Because the doublet microtubules of the axoneme are continuous with the A- and B-tubules of the basal body, sliding of the axonemal doublet microtubules causes compression and tensile forces to be experienced at the basal body

(Fisch and Dupuis-Williams, 2011; Riedel-Kruse et al., 2007; Vernon and Woolley, 2002;

Vernon and Woolley, 2004). The asymmetric beat stroke of motile cilia produce an asymmetry to the compressive and tensile forces experienced at the basal body. Because of this asymmetry, it is possible to track where compression or stretching is occurring at the basal body. This knowledge in a genetically tractable organism would be a very powerful system to model the forces that CBBs experience.

Tetrahymena as a Model Organism for Basal Body Research

The organism

Tetrahymena thermophila is a free-swimming unicellular ciliate that utilizes hundreds of motile cilia for hydrodynamic force-generation. Tetrahymena belong to the superphylum

Alveolata, which also contains the parasitic Apicomplexans and the aquatic Dinoflagellates and together compose one of the largest groups of the kingdom Protozoa (Cavalier-Smith,

1993). Tetrahymena are relatively large ovoid (20 m x 35 m) cells that contain 18-21 longitudinal rows of regularly spaced cilia (~30 per row; Figure 1.3). Each cilium is nucleated and stabilized by a conventional basal body. In addition, a single ciliated feeding structure, called an oral apparatus, contains 150 basal bodies segregated into four membranelles

(tetra - “four” hymena – “mouths”) and defines the organism’s anterior-posterior polarity.

Tetrahymena divide every three hours in a process that requires massive basal body

duplication to ensure that each daughter cell inherits an equal complement of cilia.

Tetrahymena genetics allow for the generation of genomic knock-outs, knock-ins and inducible promoter systems, and a sequenced and annotated Tetrahymena genome was recently published (Eisen et al., 2006). With sophisticated molecular genetics, defined axes of organismal polarity and a tightly controlled linear arrangement of duplicating basal bodies,

Tetrahymena is an outstanding cellular model for investigating the basic mechanisms of polarized basal body assembly and organization.

Basic Basal Body Structure

Tetrahymena basal bodies are structurally similar to basal bodies in other eukaryotes. Mature Tetrahymena basal bodies are 500-600 nm in length and 180-220 nm in diameter (Allen, 1969). The length of the basal body comprises the typical triplet microtubule blades that are arranged into a cylinder with 9-fold radial symmetry (Figure 1.4A). The

proximal end of the basal body possesses three structures that establish and maintain the cylindrical organization. First, the A- and C-tubules of adjacent triplet microtubules are connected by an A-C linkage (Figure 1.4A). Second, the proximal 60-90 nm of the basal body contains a cartwheel structure composed of a central hub and nine spokes that connect to the A-tubule of each triplet microtubule blade (Figure 1.4B). Importantly, the cartwheel is retained through the basal body lifecycle, perhaps to ensure basal body stability, as these basal bodies must resist mechanical forces from beating cilia. Third, an electron dense “collar” asymmetrically wraps around one side of the triplet microtubules

(Figure 1.4A). Distal to the cartwheel, the basal body lumen encloses an electron dense structure whose function remains poorly understood (Figure 1.4B;(Allen, 1969)). The distal end of the basal body comprises the terminal plate (the Tetrahymena transition zone), which consists of two electron dense opaque sheets that cross the lumen of the basal body (Figure

1.4B;(Allen, 1969)). While the core structure of the basal body is largely conserved,

Tetrahymena utilize a unique assemblage of accessory structures that anchor basal bodies at the cell cortex.

Additional basal body structures or accessory structures

Tetrahymena basal bodies are endowed with accessory structures to position basal bodies in coordination with the cellular polarity and to stabilize them against cilia-generated

forces (Figure 1.5). The location and composition of these structures depends on the basal body population in the Tetrahymena cell. At the cell’s anterior pole, a ring of doublet basal bodies, called dikinetids, are associated with filaments of unknown composition called the apical filament ring (Jerka-Dziadosz, 1981). Within the oral apparatus, a dense microtubule meshwork organizes approximately 150 basal bodies into its four membranelles (Williams and Frankel, 1973). For cortical basal bodies, which represent the majority of Tetrahymena basal bodies, the major accessory structures are the post-ciliary microtubules, the transverse microtubules and the kinetodesmal fiber (Figure 1.5; (Allen, 1969)). Post-ciliary microtubules nucleate from the basal body posterior face and radially project toward the posterior basal body situated in the same ciliary row. Transverse microtubules originate from the basal body anterior face and project upward and leftward (from the cell’s perspective) towards the cell cortex. The kinetodesmal fiber is a striated structure that extends from the basal body’s anterior face to the plasma membrane adjacent to the distal end of the anteriorly positioned basal body within the same ciliary row (Allen, 1969). By providing points of contact with neighboring basal bodies and the subcortical cytoskeletal network, accessory structures help establish and maintain the cellular organization and stability of basal bodies (Allen, 1969). Moreover, these structures guide the placement of newly assembled basal bodies, suggesting that cortical basal body accessory structures play an important role in cortical basal body duplication (Allen, 1969; Jerka-Dziadosz et al., 2013;

Pearson, 2014).

Basal body origins

Tetrahymena cortical basal bodies arise next to existing basal bodies in what is called centriolar basal body assembly; a daughter basal body forms orthogonally to a defined triplet microtubule at the anterior face of the proximal end of an existing mother basal body (Allen, 1969; Dippell, 1967). New assembly commences with the formation of the cartwheel and a ring of short microtubules (called a pro-basal body) that is separated from

the mother basal body by an amorphous electron-dense cloud (Allen, 1969). As the pro- basal body separates from the mother basal body, the triplet microtubules elongate and tilt towards the apical surface to dock the basal body distal end with Tetrahymena’s subcortical cytoskeletal network (Allen, 1969). The pro-basal body is positioned by the asymmetric localization of accessory structures on the mother basal body, including the kinetodesmal fiber, which ensures that the new basal body is appropriately spaced and positioned within the ciliary row (Allen, 1969). Although cortical basal bodies assemble via the centriolar pathway, the origin of oral apparatus basal bodies is unclear. One possibility is that oral apparatus basal bodies assemble de novo. Importantly, oral apparatus basal body orientation, which is random early in development, coincides with basal body linkage to an underlying microtubule network, representing a likely parallel to the process of basal body

orientation in vertebrate multi-ciliated cells (Dirksen, 1971; Kalnins and Porter, 1969;

Sorokin, 1968a; Sorokin, 1968b; Steinman, 1968; Werner et al., 2011; Williams and Frankel,

1973).

Basal body life cycle and other functions

Tetrahymena basal bodies do not function as centrioles in organizing a centrosome but rather remain docked at the cell cortex to organize cilia for the entire cell cycle. Existing mother basal bodies serve as sites of new basal body assembly that occurs throughout the cell cycle (Perlman, 1973). The nearly continuous production of basal bodies and their remarkably consistent integration into the polarized cell must be coupled with the dynamic and spatially controlled incorporation of proteins required for basal body assembly.

Basal body components

A combined use of forward and reverse genetic and proteomic approaches have identified many Tetrahymena basal body components (Galati et al., 2014; Jerka-Dziadosz et al., 1995; Kilburn et al., 2007). The triplet microtubules are comprised of canonical

andtubulin, while tubulin and  tubulin are required for basal body assembly and maintenance (Ross et al., 2013; Shang et al., 2002; Shang et al., 2005). In addition, the

Tetrahymena genome possesses  tubulin along with the ciliate specific and  tubulins, although the functions of these isoforms remain unclear (Eisen et al., 2006). Also present are the conserved UNIMOD proteins (SAS-6, CEP135/Bld10, and SAS-4/CPAP) in addition to the other strongly conserved proteins POC1 and members of the centrin family (Culver et al., 2009; Kilburn et al., 2007; Stemm-Wolf et al., 2005; Vonderfecht et al., 2012).

Notable basal body findings

Tetrahymena have played a foundational role in our understanding of basal body assembly and organization. Early studies capitalized on their polarized morphology to study the propagation and maintenance of pre-existing basal body order, which extended the

pioneering studies of Paramecium ‘structural inheritance’ by Beisson and Sonneborn into other organisms (Beisson and Sonneborn, 1965; Ng and Frankel, 1977). By mechanically inverting ciliary rows, Joseph Frankel and colleagues demonstrated that the Tetrahymena cortical architecture contains the epigenetic cues for placing new basal bodies within the polarized cell (Ng and Frankel, 1977). More recently, molecular-genetic and cytological studies identified a novel role for -tubulin in regulating basal body assembly (Shang et al.,

2005). Microtubule post-translational modifications (PTMs) are important for MT control and

Tetrahymena was fundamental in the discovery and characterization of the MEC-17/-TAT1 tubulin acetyl-transferase and the Tubulin Tyrosine Ligase Like (TTLL) modifying enzymes that glutamylate and glycylate tubulin (Janke et al., 2005; Reed et al., 2006; Wloga et al.,

2008; Wloga et al., 2009; Xia et al., 2000). Tetrahymena’s polarized cytology and ease of genetic manipulation have dramatically furthered our understanding of basal body and tubulin biology.

Strengths and future of basal body research in Tetrahymena

Coupled with new high-resolution microscopy technologies, an expanding arsenal of molecular genetic tools make Tetrahymena an immensely powerful system for the next wave of basal body research. The combined use of established forward genetics with Next

Generation sequencing enables the discovery of new molecules and mutants for further dissection of basal body assembly and organization. Moreover, use of high-resolution light and cryo-electron tomography with the numerous and easily purified basal bodies of

Tetrahymena will link the molecular and structural studies amenable to this system. The future is bright for basal body research using this evolutionarily divergent model organism to understand the most highly conserved and divergent features of basal body biology.

Conclusions and Thesis Aims

CBBs are multifunctional cellular structures. Centrioles act as the major microtubule organizing center in eukaryotic cells, and basal bodies nucleate and anchor cilia and flagella. In order to function properly CBBs must resist a significant amount of mechanical force. Centrioles experience pulling forces as they orient the mitotic spindle and basal bodies experience shear and translational forces from the beating of cilia/flagella (Abal et al., 2005; Meehl, 2016; Pearson et al., 2009b). Disruption of the ability of CBBs to resist mechanical force has dire consequences. Unstable centrioles lead to centriole fragmentation during mitosis and ultimately genomic instability, a hallmark of cancer (Basto et al., 2006; Bettencourt-Dias et al., 2005; Sir et al., 2013). Disruption of basal body stability results in loss of cilia, which causes a diverse subset of pathologies known as ciliopathies

(Badano et al., 2006). To compensate for mechanical forces, CBBs are sequentially built into a conserved structure that is highly resistant to mechanical forces (Carvalho-Santos et al., 2010). The CBB is stabilized is through domain connections between the triplet microtubules and the cartwheel, triplet microtubules and the environment and triplet microtubules with each other (Galati et al., 2014; Hiraki et al., 2007; Meehl, 2016). Another way that the CBB is stabilized is through direct stabilization of the triplet microtubules. The triple microtubules are specifically stabilized by microtubule associated proteins and post- translational modifications (Bobinnec et al., 1998b; Le Clech, 2008). The CBB remains stable despite enduring in an environment of constant mechanical forces and we are just beginning to identify the proteins and mechanisms that promote this stability.

The goal of my thesis has been to identify the factors that promote CBB stability. To do this, I have used Tetrahymena as a model system to specifically study basal body stabilization in the context of motile cilia beating. Chapter II of this thesis is published work in the journal Molecular Biology of the Cell that explores how the domain linkage protein

Bld10/Cep135 stabilizes basal bodies from the mechanical forces of ciliary beating. In

Chapter III, which is work that is in revision at the Journal of Cell Biology, I identify Fop1 as a

Bld10 and Poc1 interacting protein. Furthermore, I find that Fop1 and tubulin glutamylation stabilize the triplet microtubules. Importantly, I find that Fop1 and tubulin glutamylation are asymmetrically localized along the side of the basal body that experiences that greatest compression forces from ciliary beating. The asymmetric localization of stability proteins is the first time the radially symmetric basal body has been shown to have molecular asymmetries and this offset may represent an important mechanism for how CBB stabilization is achieved.

CHAPTER II

BLD10/CEP135 STABILIZES BASAL BODIES AGAINST CILIA-GENERATED FORCE2

Introduction

CBBs function as microtubule organizing centers in eukaryotic cells. Centrioles are part of the centrosome that organizes the interphase microtubule aster and the poles of spindles for chromosome segregation. During the G0/G1 phase of the cell cycle, the centriole converts its function from a centriole to a basal body, which organizes the microtubules of the ciliary axoneme. Cilia are cellular extensions that perform diverse roles in signaling and motility. Ciliary dysfunction causes human disorders including syndromes known as ciliopathies that exhibit a wide range of symptoms including cystic kidneys, mental retardation, microcephaly, polydactyly, respiratory illness, and retinal degeneration (Hussain et al., 2012; Kuijpers and Hoogenraad, 2011; Marszalek et al., 2000; Nachury et al., 2007;

Saeki et al., 1984). Many of the genes that cause ciliopathies encode proteins that localize to and function at the basal body (Bettencourt-Dias et al., 2011; Garcia-Gonzalo and Reiter,

2012; Nigg and Raff, 2009). Moreover, it remains to be determined whether these mutations affect ciliary function or whether they are also important in centriolar functions (Delaval et al.,

2011). Thus, understanding the mechanics for how CBBs assemble and nucleate centrosomes and cilia, respectively, is important to understand this class of human maladies.

CBBs are anchorage sites for both centrosomes and cilia. Centrioles withstand mechanical forces to facilitate spindle positioning in cells and to segregate the duplicated genome during anaphase. This is evident when cells are injected with function blocking antibodies that abrogate centriole stability causing centrosome disruption (Bobinnec et al.,

1998a). Moreover, the imbalance of microtubule generated forces causes the fragmentation

2 Portions of this chapter were previously published in Molecular Biology of the Cell, 2012 Dec;23(24):4820-32 PMID:23115304, and are included with the permission of the copyright holder.

of centrosomes (Abal et al., 2005). Consistent with a role for centrioles in withstanding mechanical forces, basal bodies resist mechanical forces from ciliary beating (Kunimoto et al., 2012). This is facilitated by the anchorage of basal bodies within the plasma membrane and the cortical architecture. Despite the important role for CBBs in withstanding mechanical forces, the molecular and structural mechanisms by which this stabilization occurs is poorly defined.

The major structural component of CBBs is microtubules. These dynamic polymers form nine modified triplet microtubule blades that are organized into a stable cylindrical structure. The stabilization of CBBs begins during new CBB formation. CBB assembly is initiated by the formation of the cartwheel. The cartwheel forms nine symmetrically spaced spokes that radiate outward from a central hub and attach to the triplet microtubules (Allen,

1969; Cavalier-Smith, 1974; Dippell, 1968; Dirksen, 1971; Sorokin, 1968a). The inner domain containing the central hub and spokes is hypothesized to establish the nine-fold symmetry (Kitagawa et al., 2011; van Breugel et al., 2011). The outer domain, which links to the inner domain, is responsible for the nucleation of the triplet microtubules (Guichard et al., 2010; Inclan and Nogales, 2001; Raynaud-Messina et al., 2004). A limited number of protein components are known to associate with these two domains. Sas6 and

Sas5/Ana2/STIL are required to assemble the inner cartwheel domain (Arquint et al., 2012;

Habedanck et al., 2005; Kitagawa et al., 2011; Stevens et al., 2010; Vulprecht et al., 2012).

Bld10/Cep135 and Poc1 are the only known outer cartwheel proteins (Hiraki et al., 2007;

Matsuura et al., 2004; Pearson et al., 2009b). Both domains are absolutely essential for the assembly of the CBB. Triplet microtubules are organized so that the microtubule minus-ends orient towards the proximal cartwheel structure and the plus-ends are positioned at a distal cap called the terminal plate or transition zone (Allen, 1969; Dippell, 1968; Dirksen, 1971;

Sorokin, 1968a). This cap containing a protein complex of CP110 and interacting proteins

ensures proper centriolar length and stabilization (Chen et al., 2002; Pearson et al., 2007;

Schmidt et al., 2009; Spektor et al., 2007; Tsang et al., 2008).

A limited number of protein components are known to stabilize these structures.

Centrobin is essential for new assembly and the stabilization of existing centrioles by promoting centriole microtubule stability (Gudi et al., 2011; Jeong et al., 2007; Zou et al.,

2005). Tetrahymena γ-tubulin loss causes the instability of basal bodies (Shang et al.,

2002). Additionally, mouse spermatocyte centriole disintegration correlates with centrin loss from the CBB and mutations in the C-terminal domain of TtCen1 causes basal body instability (Manandhar et al., 1999; Stemm-Wolf et al., 2005; Vonderfecht et al., 2011).

Function-blocking antibodies that target glutamylation modifications on centriolar tubulin disrupt centriole and centrosome stability (Abal et al., 2005). In Tetrahymena, glutamylation of microtubules is required for efficient basal body assembly (Wloga et al., 2008). These studies suggest that the stability of the CBB microtubules is important for the maintenance of these structures. The conserved CBB microtubule cylinder wall and outer cartwheel domain protein Poc1 is also required to maintain centrioles (Pearson et al., 2009b).

However, centriole assembly occurs without Poc1 suggesting that Poc1 and its localization domains on CBBs have important functions in CBB maintenance.

We searched for additional basal body stability components. In particular, we explored the possibility that outer cartwheel domain proteins stabilize CBBs. Bld10/Cep135 is a conserved outer cartwheel domain protein that is required for basal body assembly and for stable integration of Sas6 protein at basal bodies (Carvalho-Santos et al., 2010; Hiraki et al., 2007; Jerka-Dziadosz et al., 2010; Kleylein-Sohn et al., 2007; Nakazawa et al., 2007;

Roque et al., 2012). Chlamydomonas and Paramecium Bld10 makes up the tips of the cartwheel spokes (Hiraki et al., 2007; Jerka-Dziadosz et al., 2010). In addition, Drosophila

Bld10 is a microtubule associated protein that stabilizes microtubules and is required for assembly of the axoneme central doublet microtubules, suggesting that this component has

multiple roles during the basal body life cycle (Blachon et al., 2009; Carvalho-Santos et al.,

2012; Carvalho-Santos et al., 2010; Mottier-Pavie and Megraw, 2009). Because

Bld10/Cep135 is hypothesized to connect the inner cartwheel to the outer cartwheel, and binds and stabilizes microtubules, we hypothesized that, like Poc1, it also functions to stabilize the entire CBB. Tetrahymena Bld10/Cep135 (TtBld10) is required to stabilize and maintain existing basal bodies in addition to its established role in assembly of basal bodies.

We identify a novel role for Bld10/Cep135 in stabilizing existing basal bodies to resist the forces produced by ciliary beating. In summary, TtBld10 has two separable and important functions in CBB assembly and maintenance.

Results

TtBld10 is a conserved basal body cartwheel outer domain protein

The outer cartwheel domain protein Poc1 stabilizes basal bodies (Pearson et al.,

2009b). Because CBB stability is essential for its function, we searched for other outer cartwheel proteins that act as basal body stability factors. Bld10/Cep135 was a good candidate because it also localizes to the outer cartwheel domain of Chlamydomonas and

Paramecium basal bodies (Hiraki et al., 2007; Jerka-Dziadosz et al., 2010). A single

BLD10/CEP135 ortholog exists in the T. thermophila genome, and will be referred to as

TtBLD10. TtBLD10 encodes a 171 kD protein, TtBld10. As with other Bld10 family members, the protein contains extensive coiled-coil domains with two conserved regions called conserved region 1 (CR1) and conserved region 2 (CR2)(Carvalho-Santos et al., 2010;

Hodges et al., 2010)(Figure 2.1A). TtBld10 shares 42% protein sequence similarity with the human Bld10 homolog, Cep135 (Figure 2.1B and C). Consistent with a role in basal body assembly, TtBLD10 is expressed similarly to other core Tetrahymena basal body components (Miao et al., 2009).

We localized TtBld10 to determine whether TtBld10 shares a similar localization profile to that observed in other organisms. TtBld10-mCherry was expressed under the

control of its native TtBLD10 promoter in Tetrahymena cells. We found that TtBld10- mCherry localizes with TtCen1 (Stemm-Wolf et al., 2005) at all basal bodies and remains localized to basal bodies at all stages of the cell cycle (Figure 2.2A). Moreover, TtBld10- mCherry did not localize in cilia (Figure 2.2B). Similar to other organisms tested, TtBld10 is a

CBB protein.

To determine where TtBld10 localizes within the basal body architecture, we co- localized TtBld10-mCherry relative to TtCen1, which localizes asymmetrically to the proximal end and to the site of kinetodesmal fiber attachment of basal bodies (Stemm-Wolf et al.,

2005). TtBld10 localized to the proximal end of the basal body, coincident with the site of the cartwheel (Figure 2.2C) and was not found along the length of the basal body. We then localized TtBld10-mCherry relative to GFP-TtSas6a, which localizes to the central hub of the cartwheel ((Kilburn et al., 2007); Figure 2.1D). TtBld10 localizes peripherally to TtSas6a consistent with its localization to the outer cartwheel domain. Next, immuno-electron

microscopy (IEM) was employed to determine the ultrastructural localization of TtBld10- FP.

Consistent with our fluorescence data, the majority (73%) of TtBld10 immuno-gold label localized to the basal body cartwheel (Figure 2.2D). We found a small fraction (14%) of

TtBld10 localizes to the terminal plate (Figure 2.2D). Drosophila Bld10 localizes to the distal end of basal bodies and is required to form the central doublet microtubules of motile cilia

(Blachon et al., 2009; Carvalho-Santos et al., 2012; Carvalho-Santos et al., 2010; Mottier-

Pavie and Megraw, 2009). This raises the possibility that TtBld10, like DmBld10, may be required for central doublet formation. However, axoneme central doublet microtubules were normal in Ttbld10Δ cells (data not shown) suggesting that TtBld10 does not regulate the axoneme central pair microtubules in Tetrahymena as it does in Drosophila. To determine where TtBld10-GFP localizes within the cartwheel, the relative immuno-gold distribution in cross-sectional views of the cartwheel was quantified. TtBld10 associates with the ends of the cartwheel spokes (39%) and triplet microtubules (44%; Figure 2.2E). This is consistent

with Chlamydomonas and Paramecium Bld10, which also localize to the outer cartwheel and spoke tips (Hiraki et al., 2007; Jerka-Dziadosz et al., 2010). TtBld10 localization is predominantly restricted to the basal body outer cartwheel.

Ttbld10 causes the loss of basal bodies

Prior studies addressing the function of Bld10/Cep135 were limited to using hypomorphic alleles and knockdowns because a complete BLD10 genomic knockout was not accessible. Here we created, for the first time, a complete genomic knockout of BLD10

(Ttbld10Δ). Ttbld10 was induced by mating two Ttbld10 heterokaryon knockout strains to produce progeny with complete macronuclear Ttbld10 (Hai et al., 2000). Control cells were generated by mating wild-type cells with either heterokaryon knockout strain, which results in phenotypically normal cells. These control cells are referred to as TtBLD10. Ttbld10Δ cells exhibit deleterious phenotypes that are common among basal body and ciliary mutants

(Brown et al., 1999; Pearson and Winey, 2009). Ttbld10 causes cellular lethality. To determine the number of cellular divisions that Ttbld10Δ cells underwent before death, growth rates of Ttbld10Δ cell populations were quantified. Ttbld10Δ cells averaged 3.1 ± 0.7 divisions before division ceased (n=3; Figure 2.3). In addition to a reduced rate of cellular growth, the qualitative rate of cellular swimming was reduced in Ttbld10Δ cells. Moreover,

Ttbld10Δ cells exhibited a decrease in directed forward motility as seen by an increase in lateral cellular movement relative to forward movement. In summary, TtBld10 is required for cell viability, motility, and normal cell cycle progression.

Because Ttbld10Δ cells exhibited similar, albeit stronger, mutant phenotypes compared to Ttpoc1 cells, we next asked whether TtBld10 loss, like TtPoc1 loss, affects the total number of basal bodies per cell (Pearson et al., 2009b). We quantified the frequency of -TtCen1 stained basal bodies at 0, 12, 24, and 48 hours after TtBLD10 knockout. Basal body number per cell declined and organization was increasingly disrupted

with time after TtBLD10 knockout (Figure 2.4A and C). Moreover, the basal bodies of the oral apparatus disassembled in Ttbld10 cells (Figure 2.4A). To confirm that the loss of

TtBLD10 was responsible for the observed phenotypes, we rescued the Ttbld10Δ cells by reintroducing the wild-type TtBLD10 gene after knockout. Basal body number and organization were both restored by the reintroduction of TtBLD10 (Figure 2.4B). Thus,

TtBld10, like other basal body components, is required for normal basal body frequency and organization (Culver et al., 2009; Pearson et al., 2009b; Pearson and Winey, 2009; Stemm-

Wolf et al., 2005).

Bld10 is required for new basal body assembly

The inhibition of new basal body assembly causes a progressive reduction of basal bodies at each cell division. This is because basal bodies are segregated to the future cells without producing new ones to maintain the normal complement of basal bodies. In

Tetrahymena, new basal bodies form anteriorly to existing basal bodies. Basal bodies form as doublets after assembly (one new and one old). The newly assembled basal body then

moves anteriorly away from the old basal body while maturing into a basal body that nucleates a cilium (Allen, 1969; Ng and Frankel, 1977). To determine whether new basal bodies are formed in Ttbld10Δ cells, we visualized both old and new basal bodies. Old basal bodies were labeled with a marker that surrounds mature basal bodies that resemble the K-

Antigen (Williams et al., 1990), here called K-like-Antigen (Kl-Ag). This is co-localized with the pan-specific basal body marker, centrin (TtCen1) (Stemm-Wolf et al., 2005). Kl-Ag levels increase with basal body maturity (Figure 2.5A). New basal body assembly is evident as basal body doublets with TtCen1 staining but no Kl-Ag staining at the anteriorly positioned basal body (Figure 2.6A and B). Approximately 18% of the total basal bodies were newly

duplicated at both 0 and 24 hours for control cells (Figure 2.6; green arrow). The proportion of newly-assembled basal bodies dramatically decreased from 18% (0 hours) to 3% (24 hours) in Ttbld10Δ cells (Figure 2.6C). Moreover, we were unable to identify new basal body assembly in Ttbld10Δ cells at later timepoints (36 and 48 hours). We predict that the basal body assembly observed in Ttbld10Δ cells (0, 12, and 24 hours) is the result of residual, yet reduced, TtBld10 protein after knockout. The amount of new assembly decreases and is not detectable by 36 hours. Thus, TtBld10 is required for new basal body assembly.

Interestingly, we observed Kl-Ag stained foci without TtCen1 staining in Ttbld10Δ cells (Figure 2.6A; red arrowhead). The original K-Ag antibody recognizes domains within the membrane skeleton surrounding basal bodies but does not directly stain basal bodies

(Williams et al., 1990). Kl-Ag accumulates at these sites with time after basal body assembly and remains at these sites even in the absence of basal bodies in cycling cells (Figure 2.5A and Figure 2.6A; red arrowhead). Thus, loss of basal bodies does not result in the loss of Kl-

Ag staining in cycling cells. We find Kl-Ag staining in the absence of TtCen1 and these foci mark locations where basal bodies once existed and are now sites of basal body

disassembly. Furthermore, basal body disassembly occurred at immature basal bodies as judged by the reduced level of Kl-Ag staining relative to Kl-Ag levels in mature basal bodies.

This suggests that the basal bodies disassembled prior to their complete maturation.

TtBld10 is, therefore, not only required for new basal body assembly but also to stabilize developing basal bodies.

Bld10 is required to stabilize and maintain basal bodies

Basal bodies in G1-arrested cells have full levels of Kl-Ag, which indicates that they are mature. We tested whether mature basal bodies (as judged by Kl-Ag) disassemble in the absence of TtBld10. Ttbld10Δ cells were arrested in G1 so that cell division and new basal body assembly was repressed. A reduced number of basal bodies were observed in

G1 arrested Ttbld10Δ cells compared to control cells (Figure 2.7A and B). Moreover, the decrease in basal body number was time dependent, suggesting that basal bodies did not immediately disassemble but rather there was a temporal loss in basal bodies. These results further indicate that TtBld10 has an important role in maintaining and stabilizing existing basal bodies.

To directly visualize basal body disassembly, we used Kl-Ag to mark the site of basal bodies that existed prior to TtBld10 knockout. We co-localized Kl-Ag with TtCen1 in

Ttbld10Δ cells that were arrested in G1. Disassembly events (Kl-Ag staining without TtCen1 staining) in Ttbld10Δ cells were observed in a low, but significant (p<0.001), fraction of the basal body pool relative to TtBLD10 cells (Figure 2.7C and D). We hypothesize that this low fraction is due to a transient Kl-Ag signal after basal body disassembly in G1 arrested cells and this makes disassembly events difficult to capture. Moreover, the progression of Kl-Ag

disassembly was visualized in G1 arrested Ttbld10Δ cells (Figure 2.5B). Basal body and Kl-

Ag disassembly was not observed in control cells. Thus, TtBld10 is required to maintain both immature and mature basal bodies.

TtBld10 promotes triplet microtubule stability

Because TtBld10 is required for the assembly of new basal bodies and the stability of existing basal bodies, we hypothesized that TtBld10 regulates the core CBB structure. In particular, we postulated that TtBld10 regulates the triplet microtubules that comprise CBBs.

The A-tubules of triplet microtubule blades are attached to the central hub of the cartwheel

via a spoke linkage. Following assembly and attachment of the A-tubule to the cartwheels, the B- and C-tubules are then sequentially added (Dippell, 1968; Guichard et al., 2010). To determine if this organization is affected by TtBld10 loss, we visualized the basal body ultrastructure in Ttbld10Δ cells. Ttbld10Δ cells at 12 hours post knockout were prepared for transmission electron microscopy (TEM) (Dahl and Staehelin, 1989; Meehl et al., 2009;

Winey et al., 2012). 71% of the Ttbld10Δ basal bodies exhibited defects that were not found in control basal bodies (Figure 2.8; n=100 basal bodies). The Ttbld10 associated defects in triplet microtubules were categorized into three classes. 74% of the microtubule defective

Ttbld10Δ basal bodies were missing a single or multiple tubules of the microtubule triplet blade causing basal bodies to have only doublet or singlet microtubules in at least one of the basal body triplet microtubule positions (Figure 2.8B; Class 1). In cases where a doublet was present instead of a triplet the C-tubule was missing most commonly (65% of Class 1 mutants); however, a significant fraction of A-tubules were also missing (22% of Class 1 mutants). In cases where a singlet was present instead of a triplet the B-and C- tubules were always missing. In 56% of Class 1 basal bodies, the missing tubule was lost from the entire basal body length. In the remaining Class 1 samples, the basal body proximal-end contained all three tubules of the microtubule triplet and the segment distal to the cartwheel exhibited a decreased number of tubules, generating doublet or singlet morphology. The instability of tubules of the basal body triplet (Class 1) was the major defect found in

Ttbld10Δ basal bodies.

15% of defective Ttbld10Δ basal bodies were missing at least one triplet microtubule through the entire length of the basal body (Figure 2.8B; Class 2). A similar basal body phenotype was found in Paramecium cells where PtBld10a was depleted by RNAi (Jerka-

Dziadosz et al., 2010). The Tetrahymena phenotype was separated into two sub-categories.

Class 2a consists of basal bodies with triplet microtubules that conform to the missing gap thereby decreasing the basal body diameter (Figure 2.8B; Class 2a; 10% of defective basal bodies). Class 2b consists of a missing triplet microtubule that produces a gap in the 9-fold symmetry (Figure 2.8B; Class 2b; 5% of defective basal bodies). In Class 2a, the 9-fold

symmetry was likely never established and this represents a basal body assembly defect. In

Class 2b, the missing triplet microtubule likely established correct 9-fold symmetry; however, microtubule attachment and stability was disrupted.

The third class of Ttbld10 basal body defects (Class 3; 11% of defective Ttbld10 basal bodies) consisted of a combination of the first two classes where tubules of microtubule triplets and complete triplets are missing (Figure 2.8B). Thus, triplet microtubule stabilization and organization are lost in Ttbld10Δ cells.

In addition to disruption of the individual triplet microtubule structure, we find that the majority (56%) of basal bodies in Ttbld10Δ cells display triplet microtubule orientation defects. These defects are characterized by off axis positioning of entire triplet microtubule blades (Figure 2.8C). Moreover, 77% of basal bodies that possess Class 1-3 phenotypes also exhibit triplet microtubule orientation defects. These defects, in conjunction with the triplet microtubule structural defects (Classes 1-3), suggest that TtBld10 stabilizes the structure and orientation of the basal body triplet microtubules.

TtBld10 protein stably incorporates during basal body assembly and maturation

The disassembly of both immature (new, daughter) basal bodies in Ttbld10 cycling cells, and mature (old, mother) basal bodies in Ttbld10 G1-arrested cells led us to ask when TtBld10 is incorporated at basal bodies to perform its functions. We quantified the relative amounts and the timing of when TtBld10 protein incorporates during the assembly of new basal bodies and the maintenance of existing basal bodies. TtBld10-mCherry levels were variable depending on the age of the basal body. The basal body age was estimated based on distance between the daughter and mother basal bodies. Newly assembled, daughter basal bodies are closely positioned near the mother basal body while older daughter basal bodies are more physically separated from their mother basal bodies.

Mature basal bodies had increased levels of TtBld10 protein compared to immature basal

bodies (Figure 2.9A and B). At the time when the separation of mother and daughter basal bodies can be resolved newly assembled basal bodies have a mean Bld10-mCherry fluorescence intensity of approximately 40% of the mother TtBld10-mCherry fluorescent intensity. As the basal body increases in separation from its mother, an increased level of

TtBld10-mCherry was observed until a maximum protein level was reached that was equal to that of the mother basal body. Mature basal bodies did not increase or decrease in fluorescence with time suggesting that once the basal body reaches the maximum level of

TtBld10-mCherry, this level remains constant (Figure 2.9B). A similar incorporation behavior was observed using GFP-TtBld10 (Figure 2.10A and B). Thus, TtBld10 protein levels accumulate as basal bodies temporally mature.

Fluorescence recovery after photobleaching (FRAP) was used to determine whether the incorporated TtBld10 protein is stably associated with basal bodies. Single, TtBld10- mCherry labeled mature basal bodies were photobleached and live cell imaging was used to visualize the kinetics of protein redistribution and fluorescence recovery. A low level of

TtBld10-mCherry fluorescence recovery was observed indicating that TtBld10 is stably bound to the basal body (percent recovery < 5%; Figure 2.9C). When we performed longer recovery experiments of up to 10 min, we also did not observe a significant fluorescence recovery (data not shown). This supports that model that once TtBld10 protein incorporates, it remains at basal bodies.

To determine whether TtBld10 levels were recruited to maximum levels prior to ciliogenesis, we quantified the levels of TtBld10-mCherry at basal bodies possessing a cilium. TtBld10 protein is at its maximum level in ciliated basal bodies (Figure 2.9D; arrows).

This suggests that only mature basal bodies, as judged by TtBld10 levels, produce a cilium.

Because mature basal bodies disassemble in arrested Ttbld10 cells, we hypothesized that disassembly is due to ciliary beating. The instability of basal bodies without TtBld10 led us to two simplified models for TtBld10 function. First, TtBld10 is required early during new basal

body

assembly for assembly and stabilization. Second, TtBld10 stabilizes mature basal bodies to resist forces generated by beating cilia.

Decreased ciliary beating rescues basal body instability in Ttbld10 cells

TtBld10 loss causes mature basal bodies to disassemble and ciliated basal bodies have a maximum level of TtBld10 protein (Figure 2.7A and B and Figure 2.9). This suggests that TtBld10 is required to resist the forces created by ciliogenesis or ciliary beating. We assessed whether ciliary beating promotes the disassembly of basal bodies in Ttbld10Δ cells. Tetrahymena cells use cilia-dependent forces to move. To test whether Ttbld10 basal bodies disassemble as a result of the forces produced by ciliary beating, we inhibited ciliary beating in G1 cell cycle arrested Ttbld10Δ cells at 12 and 24 hours post TtBLD10 knockout by treatment with NiCl2. We found that inhibition of ciliary beating significantly

rescued basal body frequency and organization in Ttbld10 cells compared to control cells

(Figure 2.11A). These data suggest that TtBld10 stabilizes basal bodies to resist cilia- dependent forces.

In addition to inhibiting ciliary beating, we increased the physical resistance or drag force of the media that cells swim in by increasing the media viscosity with 5% polyethylene oxide (PEO). G1 cell cycle arrested Ttbld10Δ cells in 5% PEO disassemble basal bodies at a significantly greater level compared to Ttbld10Δ cells swimming in normal media (Figure

2.11B). The frequency of basal bodies was not affected in control cells treated with 5% PEO suggesting that TtBld10 is necessary to stabilize basal bodies from the increased drag force.

Discussion

Basal bodies are organizing centers for the axoneme microtubules of cilia. To resist the extreme mechanical forces subjected on them, basal bodies must stabilize the microtubules that comprise these structures. The triplet microtubule blades are important targets for basal body maintenance. We demonstrate that TtBld10 localizes to the base of the triplet microtubules at the outer cartwheel domain and that TtBld10 acts as a stability factor for basal bodies. TtBld10 is required to stabilize both immature and mature basal bodies. Our data indicate that an early incorporating population of TtBld10 protein is necessary to assemble and stabilize immature basal bodies. Subsequently, a later incorporating population of TtBld10 protects mature basal bodies against the forces of ciliary beating.

Bld10 stabilizes basal bodies

Both immature and mature basal bodies disassemble in the absence of TtBld10. We hypothesize that immature basal bodies disassemble in cells progressing through the cell cycle because they do not have a full complement of TtBld10 protein at the time of TtBLD10 knockout. In contrast, G1 arrested cells have a full complement of TtBld10 and Kl-Ag and are ciliated at the time of TtBLD10 knockout suggesting that mature basal bodies also

disassemble in Ttbld10 cells (Figure 2.7 and Figure 2.9D). However, in order to completely mature, CBBs progress through more than a single cell cycle (Nigg, 2007). At the time of starvation there is a heterogeneous population of basal bodies in each cell. Some basal bodies have progressed through one or more cell cycles and a second population of basal bodies has not progressed through a single cell cycle. Thus, the number of cell cycles that the basal bodies have progressed through may affect TtBld10-dependent basal body maintenance. This suggests that TtBld10 requires additional cell cycle progression to become competent to fulfill its stabilization duties.

Alternatively, TtBld10 may exhibit protein turnover that was not detected by our

FRAP experiments. Loss of TtBld10 protein at mature basal bodies by turnover may render

Ttbld10 basal bodies unstable. The latter possibility is unlikely because we do not observe disassembly of all basal bodies over time. Instead the level of basal body disassembly plateaus after approximately 36 hours (Figure 2.7D, data not shown). Another alternative is that basal bodies disassemble by an age-dependent turnover mechanism that is accelerated in Ttbld10 cells. Our data indicates that the disassembly of basal bodies is not caused by increased basal body turnover. If basal bodies were to disassemble as a result of normal basal body turnover, we would expect continued reduction of basal bodies to zero following

Ttbld10 knockout. Moreover, basal bodies do not disassemble in control cells that are arrested in G1 as would be expected if there was age-dependent basal body turnover

(Figure 2.7). These studies show that basal bodies are stable structures that rarely turnover or disassemble in normal Tetrahymena cells. Once assembled, basal bodies survive for many cell generations.

Bld10 is required for the triplet microtubule structure

Our electron microscopy (EM) analyses of Tetrahymena cells at 12 hours after

TtBLD10 knockout captured several classes of basal body abnormalities. These defects

range from the loss of tubules of the triplet microtubule blades (Class 1), the loss of complete triplet microtubule blades (Class 2), and the loss of triplet microtubule blade orientation (Figure 2.8). All of these phenotypes suggest that TtBld10 is important to establish and stabilize the basal body triplet microtubules. 74% of the defective

Ttbld10basal bodies were missing either the A- or C-tubules of the triplet microtubules. It is interesting to note that C. elegans centrioles lack triplet microtubules and possess a nine- fold array of singlet microtubules (Wolf et al., 1978). A BLD10/CEP135 ortholog is not found in the C. elegans genome (Carvalho-Santos et al., 2010; Hodges et al., 2010), suggesting that triplet microtubules, and in particular C-tubules, require Bld10/Cep135 for their formation and/or maintenance. Chlamydomonas - and -tubulin mutants disrupt the tubules of triplet microtubule blades (Dutcher et al., 2002; Fromherz et al., 2004). These phenotypes were specific to the B- and C-tubules and the majority (65%) of the tubules lost in Ttbld10 cells were C-tubules. Interestingly, the A-tubule was missing in 22% of the tubule-missing basal bodies (Class 1). The A-tubule loss is surprising because this tubule forms first during

CBB assembly (Dippell, 1968; Guichard et al., 2010). Subsequently, the B- and C-tubules form from the A-tubule and the B-tubule shares protofilaments with the A-tubule (Li et al.,

2012; Tilney et al., 1973). This argues that the basal body triplet microtubules formed first but that our images capture a state of A- and C-tubule disassembly. This may be an intermediate step toward basal body disassembly in the absence of TtBld10. Carvalho-

Santos and colleagues show that Drosophila Bld10 binds to and stabilizes the central pair microtubules of the ciliary axoneme (Carvalho-Santos et al., 2012). We hypothesize that cartwheel localized Tetrahymena Bld10 binds to and stabilizes the tubules of the triplet microtubule blades that make up the basal body structure.

Paramecium PtBld10a knockdown by RNAi causes the loss of entire triplet microtubule blades similar to the Ttbld10 Class 2 phenotype (Figure 2.8; (Jerka-Dziadosz

et al., 2010)). However, the loss of individual tubules within triplet microtubule blades of

Paramecium basal bodies was not observed. This is the major phenotype observed in

Tetrahymena basal bodies 12 hours after TtBLD10 knockout. This distinction between phenotypes in these two ciliate organisms may be explained by incomplete knockdown of

PtBld10 or unique functions for the second Paramecium Bld10 (PtBld10b) paralog in regulating individual microtubule stability. Like Paramecium, Chlamydomonas Bld10 stabilizes entire triplet microtubule blades and individual tubule loss was not observed in

Chlamydomonas Bld10 truncation mutants (Hiraki et al., 2007). An alternative explanation is that our ultrastructural studies captured an earlier phenotype (loss of individual tubules) after

TtBld10 loss that was not observed in the prior studies. Ultimately, all of these studies indicate that Bld10/Cep135 is necessary to stabilize the triplet microtubule blades.

The orientation of the remaining intact microtubule blades is abrogated in a majority of Ttbld10 basal bodies (Figure 2.8C). Perturbation of blade orientation may be caused by disruption of the linkage between triplet microtubule blades and the inner cartwheel and/or by disruption of the linkage between microtubule blades themselves. Bld10 comprises the spoke tip, which connects the inner cartwheel domain to the outer cartwheel domain (Figure

2.2; (Hiraki et al., 2007; Jerka-Dziadosz et al., 2010)). While we do not see shorter cartwheel spokes in Ttbld10 basal bodies, many Ttbld10basal bodies are oval shaped instead of circular when viewed as a cross section through the cartwheel (data not shown). The oval shape of Ttbld10basal bodies suggests that there is a loss of the support between the inner and the outer cartwheel. In addition, TtBld10 may function as a linker between the A- and C-tubules of adjacent triplet microtubule blades (A-C linker) that are required to connect one triplet microtubule blade to its nearest neighbor. The coiled-coil domain of TtBld10, when fully elongated, can stretch from the spoke tips to well beyond the C-tubule of an adjacent microtubule blade and it is possible that TtBld10 acts at both the A-C microtubule

linkage and the cartwheel spoke tip as a scaffold to organize the outer cartwheel domain and to link with the inner cartwheel.

In summary, our ultrastructural analyses suggest that Bld10/Cep135 has a unique function in regulating microtubule stabilization and orientation at the basal body cartwheel.

This provides the foundation for basal body stability and when defective the tubules disassemble leading to a collapse of the system.

Bld10 stabilizes basal bodies to resist cilia-generated forces

Basal bodies require TtBld10 to resist forces produced by ciliary beating (Figure

2.11A). This is evident by the rescue of G1 arrested, Ttbld10cells to normal basal body numbers by reducing ciliary beating with NiCl2 (Larsen and Satir, 1991). Thus, TtBld10, in addition to promoting new basal body assembly, is necessary to stabilize basal bodies as they anchor and counteract mechanical forces generated from beating cilia. Based on studies showing that Drosophila Bld10 binds to and stabilizes the axoneme central pair microtubules (Carvalho-Santos et al., 2012), we hypothesize that TtBld10 functions to stabilize the basal body triplet microtubules. Moreover, it remains to be determined whether

Bld10/Cep135 stabilizes centrioles in mitotic cells to render them resistant to the mechanical forces produced by the mitotic spindle.

We next tested whether increasing the drag force on cilia and, therefore, basal bodies increased the level of basal body disassembly in Ttbld10cells. By increasing the viscosity of the media that the cells swim in, basal body disassembly was increased in

Ttbld10cells but not in control cells (Figure 2.11B). The increased viscosity elevates the total drag force and load on cilia causing increased basal body disassembly. The amplified force load from increased viscosity is complicated by a decrease in ciliary beat frequency that decreases the viscous load on cilia and basal bodies. However, we postulate that increased load is still experienced by these basal bodies. Moreover, metachronal beating is

disrupted by increasing the viscosity of the medium and this may somehow contribute to basal body disassembly (Machemer, 1972). We predict that a significant force increase disrupts TtBld10-dependent basal body stabilization that is important for efficient ciliary beating that requires basal body anchorage and mechanical force resistance. In metazoans, this is important for muco-ciliary clearance where cilia of multi-ciliated epithelial cells beat in a metachronal fashion to move a highly viscous mucus layer (Hill et al., 2010). We predict that the loss of basal body resistance to such forces may contribute to respiratory illness.

In summary, Bld10/Cep135 performs two distinct roles at basal bodies. Bld10 is essential for the assembly and stabilization of basal bodies. We show here that Bld10 stabilizes basal bodies so that they can withstand the mechanical forces exerted by undulating cilia.

Materials and Methods

T. thermophila cell culture

All strains used were grown in 2% SPP media (2% protease peptone, 0.2% glucose,

0.1% yeast extract and 0.003% Fe-EDTA) to mid-log phase at 30°C, unless otherwise indicated. Cells were considered mid-log phase at a density of approximately 3x105 cells/mL as determined using a Coulter Counter Z1 (Beckman-Coulter) or a hemocytometer. To arrest cells in G1 of the cell cycle, cells were washed and resuspended in starvation media

(10 mM Tris-HCl, pH 7.4). A small fraction of cells become arrested in G2; however, the majority of cells that are the focus of this study arrest in G1.

Perturbations that affect the rate of ciliary based swimming (0.1 and 0.5 mM NiCl2, dependent on the experiment) (Larsen and Satir, 1991) or 5% poly ethylene oxide (PEO)) were introduced at the time of Ttbld10with drug selection (0 hours). Ciliary inhibition by treatment of cells with NiCl2 was visually confirmed by a decreased cellular swimming rate.

TtBLD10 identification and conservation

The Tetrahymena thermophila BLD10 gene was identified by searching the genome for the reciprocal best BLAST hit to human CEP135 and Chlamydomonas BLD10. We identified TTHERM_01164140 to fit this criterion. A Phylogenic tree was generated using a web-based phylogeny tree generation (Phylogeny.fr) (Dereeper et al., 2008). Sequences were aligned using MUSCLE (Edgar, 2004). Following alignment, gaps were removed and the phylogenetic tree was generated using the maximum likelihood method in the PhyML program (v. 3.0 aLRT) (Anisimova and Gascuel, 2006; Castresana, 2000; Guindon and

Gascuel, 2003). 100 bootstrap replicates were performed. Graphical representation was generated using TreeDyn (Chevenet et al., 2006) (Figure 2.1).

Plasmids

A TtBld10-mCherry strain was constructed by transforming cells with p4T2-

1:BLD10:mCherry. This cassette integrates at the endogenous TtBLD10 locus and remains under the control of the endogenous promoter. p4T2-1:BLD10:mCherry was generated by

PCR amplifying (5’-CGggtaccGAAGTTGATAACTGTAAGTATAC and 5’-

CGgaattcATTATTATTTTTAGATTTAGTAGAGCTTGGAGG) and cloning the final 1.5 kb of

TtBLD10 without the TGA stop codon into p4T2-1-mCherryLAP (Winey et al., 2012). A 0.9 kb fragment downstream of the TGA stop codon (5’-CGggatccTGCTCCATTCATATTTCTAT and 5’-CGgagctcAATATATCTACTCTAGCTTC) was then cloned into the plasmid to create p4T2-1:BLD10:mCherry. This plasmid contains NEO2 drug selection. A similar strain was also created using the same strategy to produce an C-terminal GFP fusion called p4T2-

1:BLD10:GFP.

The bld10 strain was generated using a BLD10 knockout cassette (pbld10::NEO2).

This construct was created by inserting 0.9 kb downstream of the TGA stop codon (5’-

CGggatccTGCTCCATTCATATTTCTAT and 5’-CGgagctcAATATATCTACTCTAGCTTC) as described above into p4T2-1-mCherryLAP to make p4T2-1-mCherryLAP:BLD10DS. A 1.3

kb fragment upstream of the ATG start codon was then PCR amplified (5’-

CGggtaccGACTTTGGACAATTTTGCTG and 5’-CGctcgagGTGATGCTAATTTGCTTCG) and cloned into p4T2-1-mCherryLAP that contains a NEO2 knockout cassette. This was cloned into a site that removes the mCherryLAP sequence but maintains the NEO2 cassette for drug selection.

To rescue the Ttbld10 strain, we generated a genomic clone of TtBLD10 with DNA flanking the BLD10 open reading frame (pBS:BLD10). A genomic TtBLD10 fragment with flanking sequence of 7.7 kb was PCR amplified (5’-CGggatccGACTTTGGACAATTTTGCTG and 5’-CGgagctcAATATATCTACTCTAGCTTC) and cloned into pBluescript (Studier and

Moffatt, 1986). The rescue cassette was then released from the vector backbone using a

Sac1 and Xho1 double digest before transformation into Tetrahymena cells.

Macronuclear transformation

GFP and mCherry fusion proteins were inserted into the macronucleus by biolistic transformation (Bruns and Cassidy-Hanley, 2000). Transformed cells were selected by using paromomycin (200 g/mL) drug to select for the NEO2 gene (Gaertig et al., 1994; Hai et al.,

2000). To increase the copy number of TtBld10-mCherry, the cells were selectively assorted by incrementally increasing the dosage of paromomycin.

Generation of the Ttbld10Δ strain

Complete genomic knockout of TtBLD10 (Ttbld10Δ) was achieved by using targeted homologous recombination to delete the germline micronuclear TtBLD10 gene using biolistic bombardment (Bruns and Cassidy-Hanley, 2000). The TtBLD10 locus was targeted using a cassette where the entire open reading frame was replaced by the NEO2 (Hai et al., 2000).

After confirming the generation of a heterozygous micronuclear Ttbld10 by genetic and

PCR-based strategies, star crosses were performed to generate two homozygous micronuclear knockout strains of different mating types (MATII and MATVI). Two confirmed

micronuclear knockout heterokaryon strains, Bld10KO1A.4 and Bld10KO 1A.4.1A, were created. Mating of Bld10KO1A.4 and Bld10KO1A.4.1A results in resistance of the progeny to paromomycin and this is coincident with the removal of TtBLD10 from the expressed macronucleus. Moreover, drug selection with paromomycin ensures that cells that did not pass through mating to knockout TtBLD10 were eliminated. For these studies, there is a small fraction of contaminating wild-type cells (approximately 5%) that do not mate and die from paromomycin treatment after approximately 24 hours post drug treatment. These cells were distinguishable from the bld10 cells and were excluded from the experiments in both fluorescence and immuno-EM studies.

BLD10 knockout

Homozygous heterokaryon strains Bld10KO1A.4 (MATVI) and Bld10KO 1A.4.1A

(MATII) that are described above were grown to mid-log phase (approximately 3x105 cells/mL) and then washed and maintained in starvation media (10mM Tris-HCl, pH.7.4) for

14 hours. Equal numbers of cells were mixed in large flasks to induce mating. 10 hours post mating initiation and equal volume of 2x SPP media was added to the mating cells (>90% mating efficiency). Paromomycin (200 g/mL) was then added 7 hours after media addition and this timepoint was the starting or 0 hour timepoint of TtBLD10 knockout.

Light microscopy

Fluorescence imaging and fluorescence recovery after photobleaching (FRAP) were performed as previously described in (Pearson et al., 2009a). In this study, a Nikon Ti

Eclipse (Nikon Instruments Inc.) inverted microscope with a Nikon 100X PlanApo NA 1.4 objective was used. Images were captured with an Andor iXon EMCCD 888E camera

(Andor Technologies). Image analysis and quantification was performed using NIS Elements imaging software (Nikon Instruments Inc., Melville, NY). All images taken were acquired at

room temperature. Acquisition times for images ranged between 50 and 500 msec, depending on the experiment.

The number of basal bodies per unit length or basal body frequency was quantified by counting the number of basal bodies (marked by TtCen1 staining) along a 10 μm segment of a ciliary row or kinety within the medial half of the Tetrahymena cell (Pearson et al., 2009b). Basal body frequency was quantified in interphase cells only to ensure that the number of basal bodies per 10 μm was constant. A minimum of 100 data points were quantified for each condition by measuring at least five ciliary rows and a minimum of 20 cells. All experiments were performed in triplicate.

Fluorescence intensities of TtBld10-mCherry at basal bodies for the maturation of

TtBld10 levels and FRAP were quantified as previously described (Pearson et al., 2009a).

Briefly, a 5 x 5 pixel region was placed over the basal body of interest and then four surrounding 5 x 5 pixel regions were used to quantify the background fluorescence levels.

The mean of the four background regions was measured and this was subtracted from the basal body fluorescence value. This corrected value was determined for each data point.

We used FRAP to quantify the dynamics of TtBld10 protein association with

Tetrahymena basal bodies using methods similarly described in (Pearson et al., 2009b).

Briefly, TtBld10-mCherry labeled basal bodies were photobleached by administering a focused 561 nm laser light pulsed on for a 45 msec exposure to bleach approximately 75% of the total basal body localized TtBld10-mCherry signal. The recovery of fluorescence was then followed in a subsequent timecourse using fluorescence imaging. The limited recovery that was observed with TtBld10-mCherry precluded us from measuring a final rate of recovery or an accurate assessment of the recovery amount (approximately 5% recovery).

The photobleaching caused by image acquisition was corrected for by quantifying the loss in fluorescence intensity of a neighboring, unbleached basal body and correcting the bleached levels. To further account for photobleaching, we also performed longer interval recovery

studies by taking the timecourse out to 5-10 min with limited image acquisition until the end of the timecourse. Under these conditions, we still did not observe additional recovery of

TtBld10-mCherry. These results suggest that TtBld10 stably incorporates and remains at the basal body.

Immunofluorescence

Immunofluorescence was performed as previously described (Cole et al., 2002).

Cells were washed once in PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, 2 mM MgCl2, pH 6.9) and then fixed in formaldehyde fixative (1% paraformaldehyde, 0.2%

Triton X-100, PHEM buffer) for one minute. Cells then were washed three times in either

0.5% boiled goat serum (BDS) or (0.1%) bovine serum albumin (BSA) in PBS (BDS-PBS or

BSA-PBS) and incubated in primary antibody in 5% BDS-PBS or 1% BSA-PBS for 24 hours at 4°C. The primary antibodies that we used were -TtCen1 (Stemm-Wolf et al., 2005); - centrin clone (20H5, Millipore)(Uzawa et al., 1995); -Kl-Antigen (10D12, (Pearson et al.,

2009a; Shang et al., 2002)); α- α-tubulin (DM1A, Sigma-Aldrich Corp., St. Louis, MO) (Blose et al., 1984). Following primary antibody incubation, cells were washed three times and then incubated in secondary antibody (Alexa Fluor 594 or 488 goat -rabbit IgG, Alexa Fluor 594, or 488 goat -mouse IgG; Invitrogen) diluted in 5% BDS-PBS or 1% BSA-PBS. After secondary incubation, cells were washed three times and adhered to poly-L-lysine-coated coverslips. Coverslips were mounted using Citifluor mounting media (Ted Pella, Inc.) and sealed using clear nail polish.

Transmission electron microscopy (TEM)

A Tetrahymena strain expressing an endogenous C-terminal TtBld10-GFP fusion was grown to mid-log phase and then prepared for immuno-electron microscopy (IEM) using high pressure freezing and freeze substitution (HPF-FS) (Dahl and Staehelin, 1989; Meehl et al., 2009). Rabbit generated -GFP antibodies were used to localize TtBld10-GFP

followed by incubation with -rabbit secondary antibodies conjugated to 15nm gold particles.

TtBld10 was then localized in 60 nm sections using transmission electron microscopy

(TEM). Images were collected using a Philips CM10 electron microscope (Philips) equipped with a Gatan BioScan2 CCD camera (Gatan).

For structural analyses of Ttbld10basal body defects, Ttbld10 and control cells were subjected to HPF-FS after 12 hours of TtBLD10 knockout. Samples were prepared as previously described (Pearson et al., 2009a). Images were acquired using an FEI Technai

G2 equipped with a Gatan Ultrascan digital camera. All images were processed for figures using Corel Draw (Corel Corporation).

CHAPTER III

MOLECULAR ASYMMETRIES STABILIZE BASAL BODIES AGAINST CILIARY

BEATING FORCES

Introduction

Motile cilia are cellular appendages that produce hydrodynamic forces for diverse functions from cellular motility to fluid flow in the respiratory tract, brain ventricles and oviduct (Lyons et al., 2006; Sawamoto et al., 2006; Wanner et al., 1996). Loss of directional fluid flow causes human disorders including primary cilia dyskinesia (PCD), hydrocephalus, and infertility (Afzelius, 1976; Afzelius and Eliasson, 1983; Greenstone et al., 1984; Ibanez-

Tallon et al., 2002). The ciliary axoneme, comprises nine doublet microtubules (A-B microtubules) arranged radially around two singlet microtubules (Silflow and Lefebvre,

2001). Cilia generate hydrodynamic force through axonemal dynein-driven sliding of doublet microtubules relative to each other (Holzbaur and Vallee, 1994). This activity leads to an asymmetric beat stroke comprising two phases: a power stroke, in which the extended cilium moves perpendicular to the cell surface; and a recovery stroke, where the bent cilium moves parallel to the cell surface thereby returning the cilium for another cycle (Chilvers and

O'Callaghan, 2000).

Basal bodies nucleate and anchor motile cilia. They comprise nine sets of radially symmetric triplet microtubules (A-B-C microtubules) organized around the cartwheel and linkages between neighboring triplet microtubules at the proximal end of the basal body. The distal end is capped by the transition zone or terminal plate. Since the A-B microtubules of basal bodies are continuous with the axoneme, basal bodies directly experience cilia- generated forces (Allen, 1969; Dippell, 1967; Dirksen, 1971). Despite the persistent exposure to ciliary force, basal body triplet microtubules are stable. They resist microtubule stressors such as cold and microtubule poisons, and they remain intact during normal ciliary beating (Abal et al., 2005; Bobinnec et al., 1998a; Brinkley and Cartwright, 1975; Kunimoto

et al., 2012). Nonetheless, defects in specific basal body components destabilize basal bodies, rendering them sensitive to cilia-generated mechanical forces (Bayless et al., 2012;

Galati et al., 2014; Schouteden et al., 2015).

The forces experienced at basal bodies are best understood for sperm flagella that undulate symmetrically along a two-dimensional axis as opposed to the three-dimensional axis of motile cilia and algal flagella (Brokaw, 1991; Brokaw et al., 1982; Riedel-Kruse et al.,

2007; Vernon and Woolley, 2004). In the sperm flagella axoneme, groups of microtubule doublets are coupled together forming two unified axonemal segments (Lindemann et al.,

1992). The doublet microtubules that compose a segment slide relative the opposing segment in a piston-like fashion (Kanous et al., 1993; Lindemann et al., 1992). Sliding between segments at the basal regions of the axoneme propagates the wave form of the ciliary beat stroke from the basal body to the tip of the axoneme (Riedel-Kruse et al., 2007;

Vernon and Woolley, 2004). Furthermore, the downward and upward movement of axoneme segments produces comparable compressive and tensile forces at opposite sides of the basal body (Vernon and Woolley, 2004). The amplitude of sliding at the flagellar axoneme base is 110 nm, suggesting that basal bodies experience compression and tensile forces along this distance (Riedel-Kruse et al., 2007). In addition, translational forces that arise from side to side movement of the axoneme are also experienced by basal bodies, although these forces are less well understood (Bayly et al., 2011). Knowledge of the flagellar beat stroke has shaped our understanding of forces experienced at basal bodies, however, less is known about how basal bodies compensate for the asymmetric force that is experienced with the asymmetric beating of motile cilia within a multi-ciliary array.

The asymmetric nature of ciliary beating causes forces at basal bodies to be asymmetrically distributed. If mathematical models of flagellar beating are interpreted within the context of asymmetric ciliary beating, then the greatest compressive forces occur at the side of the basal body where the power stroke terminates (Riedel-Kruse et al., 2007; Vernon

and Woolley, 2004). Basal bodies likely possess molecular and structural asymmetries that resist and stabilize against asymmetric forces. However, no such asymmetries have been identified.

Basal bodies, centrioles and centrosomes contain stabilizing proteins that resist the mechanical forces that these structure experience (Abal et al., 2005; Bayless et al., 2012;

Gudi et al., 2011; Manandhar et al., 1999; Pearson et al., 2009b; Ross et al., 2013; Stemm-

Wolf et al., 2013; Stemm-Wolf et al., 2005). When stability components are defective, microtubule organizing centers become functionally disrupted or they physically disassemble. In the case of disassembly, basal body loss in Tetrahymena bld10 cells can be rescued by inhibiting ciliary beating (Bayless et al., 2012). Moreover, deletion of the Poc1 basal body stability protein causes basal body triplet microtubule loss from the predicted site of greatest axoneme compression force (Meehl, 2016). Thus, in the absence of stabilizing proteins, cilia generated forces cause basal body instability. Both Bld10 and Poc1 proteins localize symmetrically around the basal body and are enriched at connections between triplet microtubules and the cartwheel or at connections between the neighboring triplet microtubules, respectively (Bayless et al., 2012; Pearson et al., 2009b). These symmetrically arranged proteins contribute to distinct mechanisms of basal body stabilization that involve forming or stabilizing linkages at unique domains of the basal body.

basal bodies are also stabilized by microtubule associated proteins and tubulin post- translational modifications (PTMs) that prevent triplet microtubule disassembly, such as the attachment of polyglutamate side chains to - and -tubulin (Magiera and Janke, 2014).

Depletion of the human microtubule associated protein, CAP350, renders centriole triplet microtubules sensitive to the depolymerizing effects of nocodazole (Le Clech, 2008).

Antibodies that target microtubule glutamylation cause centriole fragmentation, presumably by blocking the association of microtubule associated proteins or by disrupting further

glutamylation, which impairs triplet microtubule stability (Bobinnec et al., 1998a). Loss of the

Tetrahymena tubulin glutamylases, TTLL1 and TTLL9, leads to a fewer ciliary rows and defects in basal body maturation, which could result from basal body structural defects

(Wloga et al., 2008). Thus, basal body stability proteins and tubulin PTMs may contribute to basal body stability by promoting triplet microtubule stability. However, it is not known whether such stabilizing factors are asymmetrically distributed within basal bodies to resist ciliary forces.

Here, we identify Tetrahymena FGFR1 Oncogenic Partner (Fop1) as a Bld10 and

Poc1 interacting protein that is required for basal body stability. Fop1 and microtubule glutamylation is asymmetrically enriched at triplet microtubules that are predicted to experience the greatest compression forces from ciliary beating. Poc1 is necessary for normal Fop1 localization to basal bodies while both Poc1 and Fop1 levels affect basal body microtubule glutamylation. These results suggest that cooperation between distinct stability pathways creates redundancy to stabilize basal bodies against asymmetric ciliary forces.

Results

Fop1 is a basal body stability protein

Mass spectrometry was used to identify candidate Tetrahymena basal body stability proteins that interact with the known basal body stability proteins, Bld10 and Poc1

(Appendix A and B). 26 proteins with at least two spectral hits were identified, including

FGFR1 Oncogenic Partner (Fop1) (Appendix C). Fop1 (TTHERM_00537420) was identified as a candidate stability factor because its orthologs localize to centrosomes and function in microtubule anchoring (Mikolajka et al., 2006; Yan et al., 2006).

To assess whether Fop1 stabilizes basal bodies, it was depleted from Tetrahymena cells by a macronuclear gene disruption and allelic assortment, and basal bodies were monitored using immuno-fluorescence (Figure 3.1A and Figure 3.2). Tetrahymena basal

bodies are organized into longitudinal rows, so basal body frequency can be measured by determining the linear density of basal bodies within the medial half of individual ciliary rows

(Figure 3.1B). In non-dividing Fop1-depleted Tetrahymena cells, the linear density of basal bodies are reduced by 17% at 30°C (stage I; Figure 3.1C; (Williams and Scherbaum, 1959)).

Basal body loss is rescued by reintroduction of the wild-type FOP1 gene. Basal body stability mutants are often temperature sensitive and, as expected for a basal body stability protein, increased temperature (37°C) decrease the number of basal bodies in Fop1 depleted cells by 26% (Figure 3.1C; (Bayless et al., 2012; Pearson et al., 2009b)). The number of ciliary rows in all conditions remain constant suggesting that the reduced basal body frequency is not compensated for by increasing the number of ciliary rows (Figure

3.2E; (Nanney and Chow, 1974)). Decreased basal body frequency could arise from reduced basal body assembly through the cell cycle (Bayless et al., 2012). Fop1 knockdown cells were arrested in the cell cycle using starvation to halt new basal body assembly and determine whether existing basal bodies disassemble (Figure 3.1D). Fewer basal bodies are observed in arrested Fop1 knockdown cells suggesting that basal bodies disassemble upon Fop1 knockdown.

We next asked whether Fop1 depleted basal bodies display ultrastructural defects that contribute to their instability. 37% of Fop1 depleted basal bodies are reduced in diameter or have longitudinal tapering distal to the cartwheel (Figure 3.1E). Moreover, C- tubules are missing or disconnected from their associated B-tubules in more than half of the

Fop1 depleted basal bodies suggesting that Fop1 stabilizes existing basal bodies by promoting C-tubule formation and/or maintenance (Figure 3.1F).

Fop1 stabilizes basal bodies from the forces of ciliary beating

To determine whether ciliary forces contribute to basal body loss in Fop1 mutants, ciliary beating was manipulated after Fop1 knockdown cells were cell cycle arrested to follow the fate of existing basal bodies. Elevated temperature increases the Tetrahymena

ciliary beat frequency, swimming rate and, ultimately, ciliary forces at basal bodies (Goto et al., 1982; Pearson et al., 2009b). Basal body loss in Fop1 depleted cells is exacerbated with increasing temperature (Figure 3.3A). Ciliary forces can also be increased by culturing cells in viscous media containing polyethylene oxide (PEO). Fop1 depleted cells grown in media containing PEO have increased basal body loss compared to untreated Fop1 depleted cells

(Figure 3.3B). Finally, cells were treated with NiCl2 to inhibit ciliary beating. NiCl2 decreases cell swimming and completely rescue the loss of basal bodies associated with shifting Fop1 depleted cells to 37°C (Figure 3.3C). Thus, Fop1 stabilizes basal bodies against the forces derived from ciliary beating.

Fop1 localizes asymmetrically to basal bodies

The Tetrahymena ciliary power stroke moves toward the cell’s posterior leading us to

hypothesize that the basal body posterior face experiences an asymmetric compressive force (Riedel-Kruse et al., 2007). Because Fop1 stabilizes basal bodies, we asked whether

Fop1 localizes asymmetrically to reinforce basal bodies at these regions. Endogenously tagged Fop1:mCherry localizes posteriorly relative to the basal body component Cen1

(Figure 3.4A). To quantify Fop1’s localization pattern, we determined the average Fop1 localization relative to the basal body components Cen1, Sas6, and Poc1, which have known positions within the basal body architecture (Figure 3.4B-E; (Culver et al., 2009;

Pearson et al., 2009b; Stemm-Wolf et al., 2005)). Linescans of Fop1 relative to each basal body component were measured and a Gaussian fit to the averaged data was produced.

Fop1 is enriched at the basal body posterior face. Co-localization of N-terminal tagged Fop1 and C-terminal tagged Fop1 shows that the protein is not extended or arranged in a polarized orientation (Fig. 3.5A). The asymmetric localization of Fop1 to the posterior triplet microtubules was confirmed using structured illumination microscopy (SIM) and immuno- electron microscopy (Figure 3.4F-H). Surprisingly, SIM imaging reveals that Fop1 is symmetrically localized during early basal body maturation and only takes on an asymmetric

distribution once basal bodies mature (Fig. 3.5B). Early after assembly Fop1 localizes as a ring around the entire basal body cylinder but then acquires an asymmetric horse-shoe shape later in maturation. The angular displacement of the horseshoe varies but, when averaged over 45 basal bodies, the apex of the horseshoe preferentially localizes to the basal body posterior face (Fig. 3.5C). Taken together, as basal bodies mature and nucleate cilia, Fop1 accumulates at the basal body region predicted to experience the greatest compressive forces produced by ciliary beating.

Poc1 is necessary for normal incorporation of Fop1 into basal bodies

Poc1 localizes uniformly around basal body triplet microtubules; yet, in poc1 cells, the triplet microtubules at the basal body posterior face frequently disassemble (Meehl,

2016; Pearson et al., 2009b). Fop1 also localizes to the basal body posterior face suggesting that Poc1 and Fop1 stabilize this basal body domain predicted to experience compressive forces (Figure 3.4). Additionally, proteomic analyses suggest that Poc1 and

Fop1 physically interact (Appendix B). To determine whether they functionally interact, we measured the levels of endogenously tagged Fop1:mCherry in poc1cells. Fop1 levels do not change in poc1cells (Figure 3.6A). Similarly, Poc1:mCherry levels do not change upon

Fop1 depletion (Figure 3.6A). Thus, Poc1 and Fop1 are not required for each other’s basal body localization. To determine whether Poc1 and Fop1 can promote each other’s incorporation, either Poc1 or Fop1 was overexpressed and the basal body levels of the reciprocal protein was measured. Poc1 overexpression increases Fop1 basal body levels whereas Fop1 overexpression does not change Poc1 basal body levels (Figure 3.6B, C).

This suggests that Poc1 promotes Fop1 basal body incorporation, although Poc1 is not required for Fop1’s basal body localization.

To better understand how Poc1 promotes Fop1 localization, we surmised that Poc1 influences the timing of Fop1 incorporation into basal bodies to ensure that basal bodies are

stabilized at the appropriate time during basal body maturation. To test this hypothesis, we measured the initial timing and maturation of endogenously tagged protein incorporation into basal bodies. Daughter basal bodies form next to existing mother basal bodies and then separate anteriorly as they mature. Thus, the distance between mother and daughter basal bodies is a proxy for basal body maturation, and the protein fluorescence intensity ratio between the mother and daughter basal bodies is a measure of how much protein is incorporated during basal body maturation (Figure 3.6D). Fop1 incorporates early during basal body assembly and levels increase rapidly during maturation whereas Poc1 incorporates later and more slowly (Figure 3.6E, F). Both Poc1 and Fop1 reach complete protein levels before the average separation distance when basal bodies undergo ciliogenesis (2.6±0.5 m). This suggests that these stability factors incorporate prior to ciliary force generation (Figure 3.7A). Interestingly, Fop1 incorporates more slowly in poc1 cells (Figure 3.6E), and similar to the absolute levels measured above Poc1 incorporation is

unaffected by Fop1 knockdown (Figure 3.6F). In summary, although Poc1 and Fop1 do not incorporate into basal bodies at the same time, the proper timing of Fop1 incorporation requires Poc1.

Microtubule glutamylation asymmetrically localizes to basal bodies and is necessary for basal body stability

Microtubule glutamylation is asymmetrically enriched in the flagellar axoneme along the axis of the beat stroke (Fouquet et al., 1996). Furthermore, glutamylated tubulin is enriched at basal body triplet microtubules (Bobinnec et al., 1998b; Bosch Grau et al., 2013;

Suryavanshi et al., 2010; Wloga et al., 2010). This PTM has been reported to both stabilize and destabilize ciliary axonemes, while it only promotes stability at cytoplasmic microtubules and centrioles (Bobinnec et al., 1998a; Bobinnec et al., 1999; Bobinnec et al., 1998b; Wloga et al., 2010). It is unknown whether glutamylation stabilizes basal bodies against ciliary

beating forces. To address this, the basal body frequency was quantified in cells lacking the two microtubule glutamylases, TTLL1 and TTLL9 (ttll1,9, which specifically glutamylate basal body microtubules but have no measurable effect on axoneme microtubule glutamylation (Figure 3.8A, B; Figure 3.9A; (Wloga et al., 2008)). Loss of basal body microtubule glutamylation decreases the frequency of Tetrahymena basal bodies (Figure

3.8C, D). Basal body loss in ttll1,9 cells is exacerbated by elevated temperature as found in poc1 or Fop1 depleted cells, suggesting that it stabilizes basal bodies against ciliary dependent forces (Figure 3.8C-F).

We next asked whether microtubule glutamylation is asymmetrically enriched at specific basal body domains, as judged by -glutamylated tubulin antibody staining

(GT335). Longitudinally, glutamylated tubulin is found along the length of the triplet microtubules. Radially, glutamylation is concentrated at the posterior triplet microtubules

(Figure 3.8E). In mature basal bodies, microtubule glutamylation and Fop1 localize to the posterior basal body domain and are enriched where the compressive ciliary forces are predicted to be the greatest (Figure 3.8F). Microtubule glutamylation localizes internally to

Fop1 relative to the circumference of the basal body. Unlike Fop1, glutamylation at immature basal bodies remains asymmetric (Figure 3.9B). To assess the initial timing of basal body microtubule glutamylation, we determined when basal body microtubules are glutamylated relative to the incorporation of basal body stability factors (Figure 3.8H). Basal body glutamylation commences coincident with Fop1 and Bld10 incorporation, but is earlier than Poc1 incorporation (Figure 3.8H). Furthermore, the incorporation of basal body glutamylation occurs more rapidly than Poc1 (Figure 3.8H). Thus, protein factors and PTMs that stabilize basal bodies incorporate with distinct dynamics, which suggests that Fop1 and basal body microtubule glutamylation promote basal body stability through a separate mechanism than Poc1.

Poc1, Fop1 and microtubule glutamylation act in distinct pathways to stabilize basal bodies

The above studies suggest that Fop1 and Poc1 act in distinct pathways that overlap with basal body microtubule glutamylation. To determine how basal body stability factors affect basal body microtubule glutamylation, basal body glutamylation levels were quantified in poc1 and Fop1 knockdown strains. Surprisingly, loss of Poc1 or Fop1 increases basal body glutamylation, while their overexpression decreases glutamylation (Figure 3.10A, B).

This suggests that basal body glutamylation does not require Poc1 or Fop1, but instead, the increased glutamylation upon Poc1 or Fop1 loss might serve to compensate for basal body instability in their absence. Alternatively, loss of Poc1 and Fop1 may promote the accessibility of TTLL modifying enzymes to allow for additional glutamylation, facilitating microtubule severing through Katanin (Lacroix et al., 2010; Sharma et al., 2007).

Ultimately, we find that microtubule glutamylation and stability factors have an inverse relationship.

If basal body glutamylation increases to compensate for the loss of Poc1, then it would follow that decreased glutamylation in cells depleted for Poc1 would exacerbate basal body instability. To test the hypothesis that microtubule glutamylation compensates for stability factor loss and to exclude the possibility that increased glutamylation is the catalyst for basal body instability in stability factor mutants, we created cells that were null for POC1,

TTLL1, and TTLL9. Surprisingly, basal body frequency in cycling cell poc1, ttll1, ttll9 triple mutants at 30°C are increased by 15% compared to the wildtype cells and are increased by

45% when cells are shifted to 37°C (Figure 3.10C). While the frequency of basal bodies in existing ciliary rows is increased, the total number of basal bodies per cell is drastically reduced by a reduction in the number of ciliary rows (Figure 3.11A). To specifically test whether existing basal bodies are less stable in the triplet mutant, we assessed basal body frequency in arrested cells where the complication of new basal body assembly and cell division no longer occurs. Triple mutant cells exhibit a 46%, 19% and 17% decrease in basal body frequency compared to wildtype, poc1or ttll1,9cells, respectively (Figures 3.6D and 3.10D; (Pearson et al., 2009b)). Thus, basal bodies employ distinct but overlapping strategies to maintain the normal complement of basal bodies in the cell.

Discussion

Motile cilia move fluid in a single direction using an asymmetric beat pattern.

Accordingly, the basal bodies that anchor motile cilia experience asymmetric mechanical forces. We identify Fop1 as a Bld10 and Poc1 interacting protein that is required to stabilize basal bodies from cilia generated forces. Unlike the Bld10 and Poc1 stability factors, Fop1 asymmetrically localizes to the basal body domain predicted to experience the greatest cilia generated forces. Moreover, like Fop1, microtubule glutamylation stabilizes and asymmetrically localizes to basal bodies. Finally, basal body stability factors act redundantly at unique basal body structural domains to stabilize basal bodies.

Asymmetrically localized proteins stabilize basal bodies

Basal bodies are longitudinally asymmetric cylinders with a cartwheel and A-C linkages at their proximal end and the transition zone at their distal end (Bayless et al.,

2015; Pearson, 2014). However, nine sets of evenly spaced triplet microtubules impart basal bodies with a radial symmetry. Despite this, new basal body assembly occurs from a predetermined triplet microtubule at the basal body proximal end (O'Toole and Dutcher,

2014; Pearson, 2014), and basal bodies attach to the cellular cytoskeleton through structures that are asymmetric in their distribution (Bayless et al., 2015). Thus, the rotationally symmetric basal body has inherent functional and structural asymmetries.

Bld10 and Poc1 localize asymmetrically to basal body proximal ends with respect to the basal body longitudinal axis (Bayless et al., 2012; Pearson et al., 2009b). Basal body proximal ends harbor two major structural components: the cartwheel and the A-C microtubule linkers. The cartwheel organizes the basal body’s rotational nine-fold symmetry via nine spokes that attach to the triplet microtubules in a process that is facilitated by Bld10

(Carvalho-Santos et al., 2012; Hilbert et al., 2016; Hiraki et al., 2007; Kitagawa et al., 2011).

A-C microtubule linkers connect adjacent triplet microtubules to each other. The proteins that make up the A-C linkers are unknown but loss of A-C linkages likely destabilize triplet

microtubules causing them to disconnect from each other and the cartwheel (Li et al., 2012).

Since poc1 cells lose triplet microtubules with disrupted A-C linkers, it is tempting to hypothesize that Poc1 promotes the A-C triplet microtubule connections (Meehl, 2016).

Both Fop1 and microtubule glutamylation asymmetrically localize to the basal body radial axis (Figures 3.4 and 3.8E). Fop1 and microtubule glutamylation localize in a horseshoe shaped profile, effectively cupping the posterior facing triplet microtubules

(Figures 3.4 and 3.8E). To our knowledge, the asymmetric distribution of Fop1 and microtubule glutamylation are the first characterized molecular asymmetries with respect to the basal body microtubule radial axis. While both Fop1 and microtubule glutamylation are generally localized to the basal body posterior face, their precise localization appears to be dynamic (Figures 3.5C and 3.9C). We hypothesize that the dynamic variability in localization reflects the state of the ciliary beat stroke during fixation. Ciliary beating may generate sufficient mechanical strain to move basal bodies thereby altering the apparent protein localization. In summary, molecular asymmetries along the basal body’s longitudinal (Bld10,

Poc1) and radial (Fop1, tubulin glutamylation) axes enhance basal body stability.

Defining radial asymmetries of basal bodies

The mechanisms that establish molecular asymmetries within the symmetric basal body scaffold remain mysterious. One possibility is that these asymmetries are guided by intrinsic, but disordered, basal body structures that are not apparent by EM analyses. With the rapid advancement of structural microscopy methods, visualizing such previously uncharacterized structural asymmetries may not be far off. Alternatively, basal body asymmetries may be established by extrinsic factors within the environment that basal bodies mature (Pearson, 2014). One extrinsic mechanism to initiate asymmetry may be the unequal forces generated by ciliary beating, which could establish asymmetric cues for protein localization. We find that immature basal bodies have symmetrically distributed

Fop1, whereas mature basal bodies that organize cilia possess asymmetrically distributed

Fop1 (Figures 3.5B and 3.4F). The total amount of Fop1 is incorporated into the basal body prior to nucleating a cilium, so Fop1 asymmetry is either defined before ciliary beating or

Fop1 is redistributed once beating occurs. This is not true of glutamylation, which remains asymmetric regardless of basal body maturation suggesting that both early and late signals promote radial asymmetries.

Distinct and overlapping pathways to stable basal bodies

Basal body molecular asymmetries are acquired at different stages of basal body maturation (Figure 3.8H), so the relative timing of acquisition may reveal functional differences amongst the stability proteins. For example, Fop1 and microtubule glutamylation associate with the triplet microtubules and incorporate early during maturation prior to nucleation of a cilium (Figures 3.4H and 3.8H). This may allow Fop1 and microtubule glutamylation to first stabilize microtubules for basal body assembly. However, Fop1 also stabilizes basal body triplet microtubules from ciliary forces. Since Fop1 incorporates rapidly, this suggests that early basal body assembly events are critical for basal body stabilization following ciliogenesis and ciliary beating (Figure 3.3).

The incorporation of basal body stability factors that localize symmetrically along the longitudinal axis (Bld10 and Poc1) is variable during basal body maturation (Figure 3.8H).

Bld10 is required for both basal body assembly and stabilization (Bayless et al., 2012; Hiraki et al., 2007; Jerka-Dziadosz et al., 2010) and it incorporates with a biphasic loading profile.

This is consistent with a model in which the early population of Bld10 is important for basal body assembly whereas, the late population stabilizes basal bodies against ciliary forces. In support of Bld10 having multiple functions, Bld10 is essential and basal body loss in bld10cells is more severe than in the other basal body stability mutants (Figures 3.1 and

3.8; (Bayless et al., 2012; Pearson et al., 2009b)). Poc1 incorporates late and only appears to be required for stabilizing basal bodies against ciliary beating (Meehl, 2016; Pearson et

al., 2009b). Overall, asymmetric longitudinal localizing proteins have diverse assembly and maturation dynamics.

In summary, our data reveal that the radially symmetric basal body is stabilized through asymmetric localization of proteins and PTMs. These stability factors localize to specific basal body structural domains that incorporate during different stages of basal body assembly and maturation. Overall, these stability factors act in distinct, yet redundant, stabilization pathways which together allow for the powerful beating of motile cilia.

Materials and Methods

Plasmids

The Fop1:mCherry strain was generated by transforming cells with p4T2-

1:Fop1:mCherry. This cassette integrates into the endogenous Fop1 locus and remains under control of the endogenous promoter. p4T2-1:Fop1:mCherry was generated by PCR amplifying an 825 bp fragment of FOP1 immediately upstream of the TGA stop codon

(5’CGggtaccCCATTACTACTCT and 3’CGgaattcTTAATCTTCAACAT) and then cloning into p4t2-1-mCherryLAP (Winey et al., 2012). An 807 bp fragment downstream of the TGA stop codon (5’CGggatccGGATAGCTTTTTT and 3’CGgagctcTTTGATCTCACAT) was then cloned into the above plasmid intermediate to create p4T2-1:Fop1:mCherry. This plasmid contains NEO2 drug selection. The Fop1:GFP construct was generated by as described above with the p4T-1-GFPLAP plasmid (Cheeseman and Desai, 2005). The fop1 knockout cassette was created by replacing the entire open reading frame with the NEO2 resistance gene. This was achieved by cloning a 835 bp fragment of the FOP1 5’UTR

(5’CGggtaccCTATTCATCAAAA and 3’CGctcgagCTGTTCAATATGC) and a fragmant of the

FOP1 3’UTR (5’CGggatccGGATAGCTTTTTT and 3’CGgagctcTTTGATCTCACAT) into the p4t2-1 plasmid.

The Fop1 rescue construct (pBSMTTGFPFOP1) was generated by creating a genomic clone of GFP fused to FOP1 under control of an inducible MTT promoter (Shang et

al., 2002). The 1185 bp FOP1 cDNA fragment was rtPCR amplified

(5’CGtacgtaATGAGAGGATCAC and 5’CGactagtTCATTAATCTTCAA) and cloned into pBSMTTGFPgtw (provided by Doug Chalker, Washington University, St. Loius, MO) via a pENTR intermediate. This rescue cassette was transformed into Fop1 KD cells integrating at rpl29.

The overexpression constructs pBSMTTGFPSAS6a and pBSMTTGFPPOC1 were generated as described above for the pBSMTTGFPFOP1, by PCR amplifying

(5’CGggatccgATGGATAGTTTATC and 5’CGaagcttTCACTAATTTTTTG) and (5’

CGggatccgATGGCTGCGCCCTGCGCGGA and 5’

GCaagcttTCATGGTGTTGCTCTCTGCA), respectively. These genomic clones were then cloned into pBSMTTGFPCHX.

Macronuclear transformation

Biolistic transformation was used to insert p4T2-1LAP constructs, pBSMTTGFP overexpression constructs, and the fop1 knockout cassette into the macronucleus (Bruns and Cassidy-Hanley, 2000). Paromomycin (200 μg/ml) was used to select for the NEO2 gene and cycloheximide (7.5 μg/ml) was used to select for the CHX resistance (Gaertig et al., 1994; Hai et al., 2000). To increase the copy number of GFP and mCherry strains, cells were selectively assorted by incrementally increasing the dose of paromomycin.

All GFP and mCherry constructs were assorted with paromomycin until bright visualization of the fluorescent tag, after which they were maintained in that level of drug selection. To ensure equal copy number of LAP constructs in experiments where

Fop1:mCherry and Poc1:mCherry levels were assessed in the presence or absence of Poc1 or Fop1 (Figure 3.6A), respectively, strains were grown with the same concentration of paromomycin for multiple passages prior to the experiment. In experiments where Sas6,

Poc1 or Fop1 are overexpressed by induction of the MTT promoter and levels of Fop1 or

100

Poc1 are measured “WT” denotes the identical MTTGFP strains but were not induced with

CdCl2 (Figure 3.6B,C).

Fop1 knockdown

Biolistic transformation was used to insert the fop1 cassette into the macronuclear

FOP1 gene. The FOP1 locus was targeted by using a cassette where the entire open reading frame was replaced with the NEO2 gene (Hai et al., 2000). Alellic assortment of the

NEO2 gene was performed by increasing the concentration of paromomycin. Increased assortment of the knockout cassette decreases the wildtype FOP1 alleles. Knockdown was verified by genomic PCR (42% reduction) and RT-PCR (58% reduction) (Figure 3.2).

Tetrahymena thermophila cell culture

Tetrahymena strains were grown in 2% SPP media (2% protease peptone, 0.2% glucose, 0.1% yeast extract, and 0.003% Fe-EDTA) to mid-log phase at 30°C, unless otherwise indicated. Cells were considered mid-log phase at a density of 3x105 cells/ml as determined using a Coulter Counter Z1 (Beckman Coulter, Brea, CA). All ∼temperature shift experiments were performed for 24 hours. Strains were cell cycle arrested by resuspension in starvation media (10 mM Tris-HCl, pH 7.4). Perturbations that affect ciliary beating (3%

PEO or 250 M NiCl2 (Larsen and Satir, 1991)) were introduced coincident with cell cycle arrest or 24 hours after arrest, respectively. The frequency of basal bodies were quantified

24 hours after cell cycle arrest in PEO experiments and 48 hours after arrest in NiCl2 exeriments. Ciliary inhibition by treatment of cells with NiCl2 was confirmed by visualizing decreased cellular swimming.

Immunoprecipitation

Bld10:mCherryLAP or Poc1:mCherryLAP constructs were transformed into B2086

Tetrahymena cells as described above. Immunoprecipitation was performed as previously described (Cheeseman and Desai, 2005; Cheeseman et al., 2004). Briefly, 2L of confluent cells (1x106 cells/mL) were washed in PBS then resuspended in lysis buffer (50mM HEPES,

101

pH7.4; 1mM EGTA; 1mM MgCl2; 100mM KCl; 10% glycerol; 0.05% NP-40) with protease inhibitors. The whole cell solution was dropped into LiNi2 to produce drops of cell lysate. 5 g of cell drops were ground in a cryo-grinder and resuspended in 1.5x lysis buffer before sonication. Samples were centrifuged at 21,600 xg and the supernatant was collected before centrifugation at 135,000 xg. Samples were immunoprecipitated with RFP coated beads and eluted with 0.1 M Glycine pH2.6 (Cheeseman et al., 2001).

Mass spectrometry

Proteins isolated from the above immunoprecipitation were identified by using an ion trap mass spectrometer (LTQ XL; Thermo Fisher Scientific) with a reverse phase gradient over C18 resin (Phenomenex). Analysis was performed using SEQUEST software as described previously (Washburn et al., 2001).

Light microscopy

Fluorescence imaging was performed as previously described (Bayless et al., 2012).

A Nikon Ti Eclipse inverted microscope (Nikon, Melville, NY) with a Nikon 100× Plan-

Apochromant numerical aperture 1.4 objective was used. Images were captured with a

CMOS (CMOS) camera (Xyla 4.2, Andor Technology). All images were acquired using

Nikon NIS Elements imaging software. Image analysis was performed using either NIS

Elements or ImageJ. All images were acquired at room temperature. Acquisition times ranged between 50 and 500 ms, depending on the experiment. Basal body frequency was quantified as previously described (Bayless et al., 2012). Quantification of basal body frequency was performed by counting the number of basal bodies along a 10 m region along a ciliary row in the medial region of the cell. Ciliary rows around the entire circumference of the cell were quantified. All experiments utilized five measurements per cell over 20 cells. All experiments were repeated in triplicate for a total of 300 counts. All counts

102

were corrected for cell length by multiplying the number of basal bodies by the ratio of wildtype to mutant cell length.

Immunofluorescence

Immunofluorescence was performed as described previously (Bayless et al., 2012).

Cells were washed in PHEM buffer (60 mM 1,4-piperazinediethanesulfonic acid, 25 mM 4-

(2-hydroxyethyl)-1-piperazineethanesulfonic acid, 10 mM EGTA, 2 mM MgCl2, pH 6.9) then fixed in formaldehyde fixative (3.2% Paraformaldehyde, 0.2% Triton X-100, in PHEM Buffer) for 5 minutes. Cells were washed three times in 0.1% Bovine Serum Albumin (BSA) in

Phosphate Buffered Solution (PBS) or (BSA-PBS) before a 24 hour incubation at 4°C with primary antibody diluted in 1.0% BSA-PBS. Primary antibodies used in this study were -

TtCen1 (Stemm-Wolf et al., 2005), and -glutamylation (GT335; Adipogene; (Wolff et al.,

1992)). Cells were then washed three times in 0.1% BSA-PBS before incubation for two hours at 25°C in secondary antibody diluted in 1.0% BSA-PBS. Secondary antibodies used in this study were (Alexa Fluor 488, 594, or 647 goat –rabbit immunoglobulin G [IgG],

Alexa Fluor 488, 594, or 647 goat –mouse IgG; Invitrogen, Carlsbad, CA). Cells were then washed three times in 0.1% BSA-PBS and 1 l of cell pellet was added to coverslip and mounted to a slide with Citifluor mounting media (Ted Pella, Redding, CA). Samples were then sealed using clear nail polish.

Structured illumination microscopy

SIM imaging was performed using the Nikon (Nikon, Melville, NY) N-SIM system on a Ti Eclipse inverted microscope equipped with a 100× CF160 Apochromat Super-

Resolution / TIRF NA 1.49 objective. Images were captured with an Andor iXon DU897 X3

512x512 EMCCD (Andor Technology). Samples were excited using an N-SIM dual band filter cube for 488 and 561 nm excitation. All images were acquired at room temperature using NIS Elements. Image analysis was performed using NIS Elements and ImageJ.

103

Transmission electron microscopy

For immuno-EM, a C-terminal Fop1-GFP fusion under endogenous expression was prepared for IEM using high-pressure freezing and freeze substitution (HPF-FS; (Dahl and

Staehelin, 1989; Meehl et al., 2009)). Incubation in rabbit-generated -GFP antibodies, followed by incubation with -rabbit secondary antibodies conjugated to 15-nm gold particles was used to localize Fop1-GFP. Fop1 was then localized in 60-nm sections by

TEM. Images were aquired using a Philips CM10 electron microscope (Philips, Eindhoven,

Netherlands) with a Gatan BioScan2 CCD camera (Gatan, Pleasanton, CA). For EM analysis of Fop1 KD basal body structural defects, Fop1 KD cells were subjected to HPF-FS as previously described (Pearson et al., 2009b). Images were acquired using an FEI Tecnai

G2 (FEI, Hillsboro, OR) equipped with a Gatan Ultrascan digital camera. All images were processed for figures using Corel Draw (Corel, Mountain View, CA).

104

CHAPTER IV

CONCLUSIONS AND FUTURE DIRECTIONS

CBBs must be able to withstand mechanical forces to function properly. There are proteins that act to stabilize CBBs, though the mechanisms for how they do so are unclear.

The goal of my thesis has been to define and understand the factors that affect CBB stability. I used the ciliate Tetrahymena thermophila as a model system to better understand basal body stabilization in the context of motile cilia generated forces. This system has proven invaluable for my research. Specifically, the robust number of basal bodies and the genetic tools to knockout a gene at a single timepoint gives a large advantage in identifying and studying basal body stability mutants. Additionally, the ability to fluorescently tag proteins has allowed me to visualize where multiple proteins localize in live cells. Using this system I have made two novel advancements in the field of CBB stabilization. First, I find that basal body stability factors stabilize against the forces of ciliary beating, second I find that stability factors can alter their localization to compensate for the areas that experience mechanical force.

The first major conclusion from my research is that basal body stability factors stabilize against ciliary generated mechanical forces. I have shown that the assembly protein Bld10 also functions as a basal body stability factor (Figure 2.4). Bld10/Cep135 is a major component of the CBB. Before my findings, it was known that Bld10/Cep135 connected the cartwheel to the triplet microtubules and was required for new CBB assembly, but a role in CBB maintenance was not known (Carvalho-Santos et al., 2012;

Hiraki et al., 2007; Jerka-Dziadosz et al., 2010). I next developed assays in Tetrahymena to manipulate the force that basal bodies experience by modulating the ciliary beat stroke. To this end, I used NiCl2 to inhibit ciliary beating and find that in the presence of reduced ciliary forces Bld10 associated loss of basal bodies is rescued (Figure 2.11). Alternatively, I used

PEO to increase the media viscosity that cilia beat through and correspondingly find that I

105

basal body loss is exacerbated compared to bld10 cells not treated with PEO (Figure

2.11). Using these force manipulation strategies, I also identified that the CBB protein Fop1 is a basal body stability protein and that it stabilizes the basal body from the forces of ciliary beating (Figures 3.1 and 3.3). Together, these results verify, that basal bodies experience force from ciliary beating. Furthermore, these experiments demonstrate that there are proteins that function by helping the basal body resist these mechanical forces.

The second major insight from my research is that stability factors localize asymmetrically at the basal body (Figures 3.4 and 3.8). This includes the stability protein

Fop1 and the tubulin post-translational modification glutmalyation, which are distinct from other basal body stability factors because they localize along the triplet microtubules. The basal body is radially symmetric, and even though it has been studied for over 60 years it has never been shown to have molecular asymmetries with respect to its radial axis. This work has taken predictions from modeling studies of flagellar basal bodies and tested them biologically (Riedel-Kruse et al., 2007; Vernon and Woolley, 2004). Importantly, the localization of stability factor asymmetry is consistent with the site of greatest compressive force and the site of initial destruction of the basal body in stability factor loss (Meehl, 2016).

Hopefully, the identification of asymmetrically localizing basal body stability factors will generate more discussion of the forces that CBBs experience.

My research leaves a few unanswered questions about CBB stabilization. One aspect of CBB stabilization that could use more examination is how the CBB matures through the addition of proteins. Assembly of the CBB is a stepwise process that relies on the correct incorporation of proteins to build a completely stable structure. The characterization of when proteins incorporate into the CBB is an area of research that has not been explored after initial assembly events. Furthermore, all of the stability factors that I have identified incorporate into the CBB after early assembly during maturation. This maturation phase of CBB development is not well understood. I have developed a novel way

106

to measure the timing and rate of protein maturation into the CBB (Figure 3.6). Using this approach, I have found that knockout of the stability factors can alter the incorporation dynamics of other stability factors, which highlights how important it is to build the CBB correctly (Figure 3.6). I can envision this assay being used to characterize the incorporation profiles of known CBB components to bring about a greater understanding of the temporal ordering of CBB assembly.

One major unanswered question left by my research is what drives asymmetric localization of basal body stability factors. The most intriguing answer is that stability factors, like Fop1, change their localization to compensate for force. However, in our preliminary data we do not see any experimental evidence for this. In newly assembled basal bodies

Fop1 is still asymmetrically localized well before ciliation takes place (Figure 3.5). There are other ways to test whether force dictates molecular asymmetries. One way to test whether force dictates the positioning of basal body stability factors is to alter the amount or directionality of force and see whether stability factors can reposition in response. The easiest way to do this in my current system would be to use NiCl2 to inhibit ciliary beating and see whether Fop1 still localizes asymmetrically. Another way to do this would be to put

Tetrahymena in a flow chamber and move fluid flow laterally across the organism then determine whether Fop1 localization changes to compensate. Another possibility is to take advantage of the ciliate Paramecium’s ability to reverse its power stroke and thus reverse the distribution of forces, though it would first have to be verified that Paramecium Fop1 is asymmetrically distributed in the first place.

Redistribution of stability factors to compensate for force cannot be the only mechanism of asymmetric localization because tubulin glutamylation is positioned asymmetrically at the basal body well before a cilium is nucleated (Figure 3.9). It is possible that the basal body experiences some level of mechanical force before ciliation due to the translational movement of the mature basal body. Because new basal bodies are templated

107

and attached to the mature basal body, any movement of the mature basal body could be experienced by the newly forming daughter basal body. Whether this force is asymmetric remains to be determined. An alternative explanation of how asymmetric modification of tubulin by glutamylation is initiated could be due to the positioning of the nascent basal body itself during early assembly. Since new basal bodies are built orthogonally off of a mature basal body it is possible that the closer localization of one face of the basal body to the mature basal body in some way predisposes that section of the basal body to receive differential amounts of protein or post-translational modifications. Although, it is still unknown whether the position of the nascent basal body during its construction is in any way deterministic of its final orientation.

Basal bodies of motile cilia do not only experience asymmetric compressive forces, they also experience an equal amount of asymmetric tensile or stretching forces. I have identified two factors that localize asymmetrically to the site of compressive forces but it remains to be determined whether there are stability factors that asymmetrically localize to the site of tensile forces. One potential candidate for this type of asymmetric localization is the post-translational modification glycylation. Tubulin glycylation occurs off of the same residues as glutamylation so in essence they compete for the same modification sites

(Wloga and Gaertig, 2010). Since tubulin glutamylation and glycylation compete for the same modification sites it stands to reason that the side of the basal body that is low in glutamylation modifications would be enriched for glycylation. Also, in the cilium glycylation functions differentially than glutamylation and their balance is essential for ciliary beating

(Bosch Grau et al., 2013). It is possible that the same can be true of the basal body since each side of the basal body has a different need for stabilization due to the differential forces experienced. It may be that there are no stability factors that asymmetrically localize to the side of the basal body that experiences tensile forces. The absence of stability factors could represent an intrinsic stability of the basal body against tensile forces. This could be

108

mediated by the transition zone through a break between the A- and B-tubules that is not easy to capture through electron microscopy. If this were the case then stretching forces would not be translated to the basal body. Even if this break were present at the site of compression, compression forces translated to the basal body. Overall, the stabilization against tensile forces remains as an intriguing question moving forward.

In conclusion, the data presented in this thesis advances the field of CBB stabilization by identification of stability factors and the novel finding that molecular asymmetries exist within the radial axis of basal bodies. Though there needs to be more work done to determine the signals guiding origination of molecular asymmetries and the nature of tensile force stabilization, the future of CBB stabilization research is promising.

109

APPENDIX A

Bld10 Immunoprecipitation

Spectral Bld10 IP Sequence Coverage Protein Name Hits H/ACA ribonucleoprotein TTHERM_000666223 47 76.6% complex protein, putative TTHERM_01164140 445 49.1% Bld10 TTHERM_00194520 49 41.4% hypothetical protein Gar1/Naf1 RNA-binding TTHERM_00929590 11 40.2% region protein TTHERM_01014750 44 31.8% chaperone protein DnaK SCF ubiquitin ligase TTHERM_00426320 7 31.3% complex protein aldo/keto reductase family TTHERM_00338200 79 27.7% oxidoreductase aldo/keto reductase family TTHERM_01002600 25 24.1% oxidoreductase pyruvate dehydrogenase TTHERM_00530750 21 19.9% complex dihydrolipoamide acetyltransferase pre-RNA processing TTHERM_00752200 18 18.9% PIH1/Nop17 protein TTHERM_00558440 41 18.4% heat shock 70 kDa protein L-isoaspartate O- TTHERM_00283890 9 16.8% methyltransferase TTHERM_000522989 2 16.7% hypothetical protein TTHERM_00125640 44 16.4% heat shock 70 kDa protein TTHERM_00471510 4 15.9% SNF7 family protein EF-hand calcium-binding TTHERM_001232262 3 15.7% domain protein TTHERM_00836580 20 14.9% tubulin TTHERM_00348510 20 14.9% tubulin TTHERM_00283340 9 14.8% hypothetical protein elongation factor Tu TTHERM_00151810 17 14.3% protein TTHERM_00190950 4 14.1% macronuclear actin TTHERM_000584709 7 13.9% kinase domain protein 25 kDa calcium-binding TTHERM_00068170 8 13.7% protein glyceraldehyde-3- TTHERM_00551160 6 13.5% phosphate dehydrogenase TTHERM_00471950 10 13.2% hypothetical protein TTHERM_00444510 7 13.0% ribosomal protein L19

110

Bld10 Immunoprecipitation Continued

Spectral Bld10 IP Sequence Coverage Protein Name Hits ATP synthase F1, beta TTHERM_00585260 5 12.7% subunit aldo/keto reductase family TTHERM_00300640 3 11.8% oxidoreductase TTHERM_00137990 8 11.7% oxidoreductase, putative cyclin-dependent kinase- TTHERM_01207660 9 11.7% like Serine/Threonine kinase family protein TTHERM_00537420 2 10.9% Fop1 hydrogenosomal ATP/ADP TTHERM_00037640 4 9.3% carrier protein heat shock-binding protein TTHERM_00171850 10 9.0% 70, ER luminal protein tRNA pseudouridine TTHERM_00095540 7 9.0% synthase elongation factor 2, TTHERM_00938820 12 8.7% putative TTHERM_00196370 7 8.7% chaperonin CPN60-1 TTHERM_00379000 11 8.1% hypothetical protein SIR2 family transcriptional TTHERM_00112480 4 8.1% regulator TTHERM_00158520 8 7.5% predicted protein pyruvate kinase complex TTHERM_01016110 6 6.3% alpha subunit TTHERM_000561429 6 5.7% MIZ zinc finger protein tubulin/FtsZ family, GTPase TTHERM_00558620 10 5.6% domain protein TTHERM_00077140 3 5.0% hypothetical protein PX-SNX8-Mvp1p-like TTHERM_00502220 3 4.9% protein transitional endoplasmic TTHERM_00365340 4 4.6% reticulum ATPase, putative acyl-CoA thioester TTHERM_00371200 4 4.5% hydrolase TTHERM_01308010 2 4.3% Poc1

TTHERM_00537180 2 4.1% hypothetical protein

TTHERM_00926980 4 3.8% acetyl-CoA acyltransferase

TTHERM_00637010 5 3.8% hypothetical protein

111

Bld10 Immunoprecipitation Continued

Spectral Bld10 IP Sequence Coverage Protein Name Hits spindle assembly TTHERM_00388200 2 3.3% abnormal-like protein, putative TTHERM_00444670 4 3.0% heat shock protein HSP90 TTHERM_00024480 2 2.9% PX domain protein TTHERM_00316230 3 2.8% tubulin glycylase 3C condensin complex subunit TTHERM_00554600 5 2.8% 2 TTHERM_00155290 4 2.7% hypothetical protein kinesin motor catalytic TTHERM_00444350 2 1.2% domain protein

112

APPENDIX B

Poc1 Immunoprecipitation

Spectral Poc1 IP Sequence Coverage Protein Name Hits H/ACA ribonucleoprotein TTHERM_000666223 23 56.2% complex protein, putative TTHERM_000522989 29 42.3% hypothetical protein TTHERM_00194520 25 30.3% hypothetical protein TTHERM_01308010 143 27.6% Poc1 pre-RNA processing TTHERM_00752200 23 27.6% PIH1/Nop17 protein Gar1/Naf1 RNA-binding TTHERM_00929590 4 27.0% region protein TTHERM_01014750 31 26.1% chaperone protein DnaK TTHERM_00471950 39 22.0% hypothetical protein ubiquitin-conjugating TTHERM_00532840 3 20.4% enzyme elongation factor Tu TTHERM_00151810 18 19.9% protein TBP-interacting DNA TTHERM_00046920 14 18.9% helicase aldo/keto reductase TTHERM_00338200 23 17.6% family oxidoreductase 25 kDa calcium-binding TTHERM_00068170 13 16.7% protein

TTHERM_00196370 15 16.6% chaperonin CPN60-1

TTHERM_00537420 5 16.5% Fop1

TTHERM_00558440 27 16.4% heat shock 70 kDa protein

TTHERM_000584709 12 16.0% kinase domain protein

hydrogenosomal TTHERM_00037640 8 15.4% ATP/ADP carrier protein aldo/keto reductase TTHERM_01002600 17 14.4% family oxidoreductase

TTHERM_00190950 13 12.8% macronuclear actin

ATP synthase F1, beta TTHERM_00585260 10 12.7% subunit

113

Poc1 Immunoprecipitation Continued

Spectral Poc1 IP Sequence Coverage Protein Name Hits elongation factor 2, TTHERM_00938820 15 11.3% putative TTHERM_00055970 3 10.3% SNF7 family protein TTHERM_00379000 17 10.2% hypothetical protein TCP-1 (CTT or eukaryotic TTHERM_00239290 8 10.2% type II) chaperonin family, gamma subunit Bmh1 14-3-3 family TTHERM_00216010 4 10.2% protein TTHERM_00283340 4 10.1% hypothetical protein 3-hydroxyacyl-CoA TTHERM_00666640 2 10.1% dehydrogenase TTHERM_00522050 5 9.8% ankyrin domain protein tyrosine 3- monooxygenase/tryptoph TTHERM_00160770 2 9.6% an 5-monooxygenase activation protein, epsilon protein TTHERM_00471510 4 9.6% SNF7 family protein GTP-binding nuclear TTHERM_00023980 2 9.3% protein Ran-1 TTHERM_00836580 18 8.8% tubulin TTHERM_00348510 18 8.8% tubulin

TTHERM_00158520 13 8.4% predicted protein G-quartet DNA-binding TTHERM_00499440 3 8.3% protein, putative TTHERM_00086760 4 8.2% co-chaperone GrpE SIR2 family TTHERM_00112480 2 8.1% transcriptional regulator NAD-dependent isocitrate TTHERM_00723390 5 8.1% dehydrogenase 2-oxo acid TTHERM_00794540 2 8.0% dehydrogenase acyltransferase pyruvate dehydrogenase TTHERM_000784559 2 7.5% E1 beta subunit heat shock-binding TTHERM_00171850 6 7.4% protein 70, ER luminal protein ADP/ATP transporter on TTHERM_00363210 2 6.9% adenylate translocase ADP/ATP transporter on TTHERM_00052310 3 6.8% adenylate translocase

114

Poc1 Immunoprecipitation continued

Spectral Poc1 IP Sequence Coverage Protein Name Hits NAD-dependent isocitrate TTHERM_00344030 12 6.5% dehydrogenase transmembrane protein, TTHERM_00127260 2 6.3% putative TTHERM_00391300 2 6.2% chaperone protein DnaJ TTHERM_00571650 4 6.2% hypothetical protein condensin complex TTHERM_00554600 8 6.0% subunit 2 eukaryotic aspartyl TTHERM_00462870 2 5.6% protease M16 family peptidase, TTHERM_00836690 2 5.6% putative tubulin/FtsZ family, TTHERM_00558620 4 5.6% GTPase domain protein 26S proteasome TTHERM_00469100 5 5.6% regulatory ATPase subunit 10B TTHERM_00137990 3 5.6% oxidoreductase, putative TTHERM_00275740 10 5.6% clathrin heavy chain acetyl-CoA TTHERM_00926980 4 5.5% acyltransferase pyruvate kinase complex TTHERM_01016110 4 5.5% alpha subunit 26S protease regulatory TTHERM_00068110 3 5.2% subunit 6B TTHERM_00317440 3 5.1% acyl carrier reductase TCP-1 (CTT or eukaryotic TTHERM_00149340 2 5.0% type II) chaperonin family, beta subunit TTHERM_00426310 4 5.0% chaperone DnaJ translation elongation TTHERM_00655820 2 4.8% factor EF-1alpha ATP synthase F1, alpha TTHERM_00571860 3 4.2% subunit TTHERM_00155290 4 4.1% hypothetical protein RPAP3 monad-binding TTHERM_00283980 2 4.0% domain protein, putative TTHERM_00011900 2 3.9% hypothetical protein

TTHERM_00690030 4 3.7% ubiquitin family protein

TTHERM_00579210 6 3.7% hypothetical protein

115

Poc1 Immunoprecipitation Continued

Spectral Poc1 IP Sequence Coverage Protein Name Hits PX-SNX-like domain TTHERM_00149760 2 3.2% protein TTHERM_00420790 3 3.2% EF-hand pair protein tesmin TSO1-like CXC TTHERM_00388590 2 3.0% domain protein DNA replication licensing TTHERM_00448570 3 3.0% factor MCM4 TTHERM_00444670 3 3.0% heat shock protein HSP90 condensin complex TTHERM_01299730 2 2.9% subunit 2 TTHERM_000242219 2 2.8% hypothetical protein TTHERM_00470990 2 2.5% DnaK protein sarco/endoplasmic reticulum calcium- TTHERM_01084370 2 2.5% translocating P-type ATPase TTHERM_00813040 2 2.4% PCI-domain protein TTHERM_000561429 3 2.3% MIZ zinc finger protein RNA recognition motif TTHERM_00051730 2 2.2% protein structural maintenance of TTHERM_00446400 2 2.1% chromosomes protein TTHERM_00637010 3 1.7% hypothetical protein TTHERM_00335640 3 1.3% hypothetical protein outer arm dynein beta TTHERM_00499300 4 1.1% heavy chain dynein heavy chain, TTHERM_00046310 4 1.1% cytoplasmic protein cation channel family TTHERM_000145989 2 0.5% transporter, putative

116

APPENDIX C

Common Proteins from Bld10 and Poc1 IP

Common from Bld10 and Protein Name Poc1 IP

TTHERM_00037640 hydrogenosomal ATP/ADP carrier protein

TTHERM_00112480 SIR2 family transcriptional regulator TTHERM_00155290 hypothetical protein TTHERM_00158520 predicted protein

TTHERM_00171850 heat shock-binding protein 70, ER luminal protein

TTHERM_00190950 macronuclear actin TTHERM_00194520 hypothetical protein TTHERM_00196370 chaperonin CPN60-1 TTHERM_00283340 hypothetical protein

TTHERM_00338200 aldo/keto reductase family oxidoreductase

TTHERM_00348510 Tubulin TTHERM_00379000 hypothetical protein TTHERM_00444670 heat shock protein HSP90 TTHERM_00471950 hypothetical protein pyruvate dehydrogenase complex dihydrolipoamide TTHERM_00530750 acetyltransferase TTHERM_00537420 Fop1 TTHERM_00554600 condensin complex subunit 2 TTHERM_00558440 heat shock 70 kDa protein

TTHERM_00558620 tubulin/FtsZ family, GTPase domain protein

TTHERM_00561680 ribosomal protein L35 TTHERM_00585260 ATP synthase F1, beta subunit TTHERM_00706300 ribosomal protein S9

TTHERM_00752200 pre-RNA processing PIH1/Nop17 protein

TTHERM_00836580 Tubulin

TTHERM_00938820 elongation factor 2, putative

TTHERM_01308010 Poc1

117

REFERENCES

Abal, M., G. Keryer, and M. Bornens. 2005. Centrioles resist forces applied on centrosomes during G2/M transition. Biology of the cell / under the auspices of the European Cell Biology Organization. 97:425-434.

AbouAlaiwi, W.A., M. Takahashi, B.R. Mell, T.J. Jones, S. Ratnam, R.J. Kolb, and S.M. Nauli. 2009. Ciliary polycystin-2 is a mechanosensitive calcium channel involved in nitric oxide signaling cascades. Circulation research. 104:860-869.

Afzelius, B. 1959. Electron microscopy of the sperm tail; results obtained with a new fixative. The Journal of biophysical and biochemical cytology. 5:269-278.

Afzelius, B.A. 1976. A human syndrome caused by immotile cilia. Science. 193:317-319.

Afzelius, B.A., and R. Eliasson. 1983. Male and female infertility problems in the immotile- cilia syndrome. European journal of respiratory diseases. Supplement. 127:144-147.

Al Jord, A., A.I. Lemaitre, N. Delgehyr, M. Faucourt, N. Spassky, and A. Meunier. 2014. Centriole amplification by mother and daughter centrioles differs in multiciliated cells. Nature. 516:104-107.

Allen, R.D. 1969. The morphogenesis of basal bodies and accessory structures of the cortex of the ciliated protozoan Tetrahymena pyriformis. The Journal of cell biology. 40:716- 733.

Alvey, P.L. 1986. Do adult centrioles contain cartwheels and lie at right angles to each other? Cell biology international reports. 10:589-598.

Anderson, R.G., and R.M. Brenner. 1971. The formation of basal bodies (centrioles) in the Rhesus monkey oviduct. The Journal of cell biology. 50:10-34.

Anisimova, M., and O. Gascuel. 2006. Approximate likelihood-ratio test for branches: A fast, accurate, and powerful alternative. Systematic biology. 55:539-552.

Arquint, C., A.M. Gabryjonczyk, S. Imseng, R. Bohm, E. Sauer, S. Hiller, E.A. Nigg, and T. Maier. 2015. STIL binding to Polo-box 3 of PLK4 regulates centriole duplication. eLife. 4.

Arquint, C., K.F. Sonnen, Y.D. Stierhof, and E.A. Nigg. 2012. Cell-cycle-regulated expression of STIL controls centriole number in human cells. Journal of cell science. 125:1342-1352.

Badano, J.L., N. Mitsuma, P.L. Beales, and N. Katsanis. 2006. The ciliopathies: an emerging class of human genetic disorders. Annual review of genomics and human genetics. 7:125-148.

Bannai, H., M. Yoshimura, K. Takahashi, and C. Shingyoji. 2000. Calcium regulation of microtubule sliding in reactivated sea urchin sperm flagella. Journal of cell science. 113 ( Pt 5):831-839.

118

Bastians, H. 2015. Causes of Chromosomal Instability. Recent results in cancer research. Fortschritte der Krebsforschung. Progres dans les recherches sur le cancer. 200:95- 113.

Basto, R., J. Lau, T. Vinogradova, A. Gardiol, C.G. Woods, A. Khodjakov, and J.W. Raff. 2006. Flies without centrioles. Cell. 125:1375-1386.

Bayless, B.A., D.F. Galati, and C.G. Pearson. 2015. Tetrahymena basal bodies. Cilia. 5:1.

Bayless, B.A., T.H. Giddings, Jr., M. Winey, and C.G. Pearson. 2012. Bld10/Cep135 stabilizes basal bodies to resist cilia-generated forces. Molecular biology of the cell. 23:4820-4832.

Bayly, P.V., B.L. Lewis, E.C. Ranz, R.J. Okamoto, R.B. Pless, and S.K. Dutcher. 2011. Propulsive forces on the flagellum during locomotion of Chlamydomonas reinhardtii. Biophysical journal. 100:2716-2725.

Beisson, J., and T.M. Sonneborn. 1965. Cytoplasmic Inheritance of the Organization of the Cell Cortex in Paramecium Aurelia. Proceedings of the National Academy of Sciences of the United States of America. 53:275-282.

Bettencourt-Dias, M., F. Hildebrandt, D. Pellman, G. Woods, and S.A. Godinho. 2011. Centrosomes and cilia in human disease. Trends in genetics : TIG. 27:307-315.

Bettencourt-Dias, M., A. Rodrigues-Martins, L. Carpenter, M. Riparbelli, L. Lehmann, M.K. Gatt, N. Carmo, F. Balloux, G. Callaini, and D.M. Glover. 2005. SAK/PLK4 is required for centriole duplication and flagella development. Current biology : CB. 15:2199-2207.

Blachon, S., X. Cai, K.A. Roberts, K. Yang, A. Polyanovsky, A. Church, and T. Avidor-Reiss. 2009. A proximal centriole-like structure is present in Drosophila spermatids and can serve as a model to study centriole duplication. Genetics. 182:133-144.

Black, M.M., and P. Keyser. 1987. Acetylation of alpha-tubulin in cultured neurons and the induction of alpha-tubulin acetylation in PC12 cells by treatment with nerve growth factor. The Journal of neuroscience : the official journal of the Society for Neuroscience. 7:1833-1842.

Blose, S.H., D.I. Meltzer, and J.R. Feramisco. 1984. 10-nm filaments are induced to collapse in living cells microinjected with monoclonal and polyclonal antibodies against tubulin. The Journal of cell biology. 98:847-858.

Bobinnec, Y., A. Khodjakov, L.M. Mir, C.L. Rieder, B. Edde, and M. Bornens. 1998a. Centriole disassembly in vivo and its effect on centrosome structure and function in vertebrate cells. The Journal of cell biology. 143:1575-1589.

Bobinnec, Y., C. Marcaillou, and A. Debec. 1999. Microtubule polyglutamylation in Drosophila melanogaster brain and testis. European journal of cell biology. 78:671- 674.

Bobinnec, Y., M. Moudjou, J.P. Fouquet, E. Desbruyeres, B. Edde, and M. Bornens. 1998b. Glutamylation of centriole and cytoplasmic tubulin in proliferating non-neuronal cells. Cell motility and the cytoskeleton. 39:223-232.

119

Bosch Grau, M., G. Gonzalez Curto, C. Rocha, M.M. Magiera, P. Marques Sousa, T. Giordano, N. Spassky, and C. Janke. 2013. Tubulin glycylases and glutamylases have distinct functions in stabilization and motility of ependymal cilia. The Journal of cell biology. 202:441-451.

Bre, M.H., B. de Nechaud, A. Wolff, and A. Fleury. 1994. Glutamylated tubulin probed in ciliates with the monoclonal antibody GT335. Cell motility and the cytoskeleton. 27:337-349.

Bre, M.H., V. Redeker, M. Quibell, J. Darmanaden-Delorme, C. Bressac, J. Cosson, P. Huitorel, J.M. Schmitter, J. Rossler, T. Johnson, A. Adoutte, and N. Levilliers. 1996. Axonemal tubulin polyglycylation probed with two monoclonal antibodies: widespread evolutionary distribution, appearance during spermatozoan maturation and possible function in motility. Journal of cell science. 109 ( Pt 4):727-738.

Brinkley, B.R., and J. Cartwright, Jr. 1975. Cold-labile and cold-stable microtubules in the mitotic spindle of mammalian cells. Annals of the New York Academy of Sciences. 253:428-439.

Brokaw, C.J. 1991. Microtubule sliding in swimming sperm flagella: direct and indirect measurements on sea urchin and tunicate spermatozoa. The Journal of cell biology. 114:1201-1215.

Brokaw, C.J., and R. Kamiya. 1987. Bending patterns of Chlamydomonas flagella: IV. Mutants with defects in inner and outer dynein arms indicate differences in dynein arm function. Cell motility and the cytoskeleton. 8:68-75.

Brokaw, C.J., D.J. Luck, and B. Huang. 1982. Analysis of the movement of Chlamydomonas flagella:" the function of the radial-spoke system is revealed by comparison of wild- type and mutant flagella. The Journal of cell biology. 92:722-732.

Brown, J.M., C. Marsala, R. Kosoy, and J. Gaertig. 1999. Kinesin-II is preferentially targeted to assembling cilia and is required for ciliogenesis and normal cytokinesis in Tetrahymena. Molecular biology of the cell. 10:3081-3096.

Bruns, P.J., and D. Cassidy-Hanley. 2000. Biolistic transformation of macro- and micronuclei. Methods in cell biology. 62:501-512.

Carvalho-Santos, Z., J. Azimzadeh, J.B. Pereira-Leal, and M. Bettencourt-Dias. 2011. Evolution: Tracing the origins of centrioles, cilia, and flagella. The Journal of cell biology. 194:165-175.

Carvalho-Santos, Z., P. Machado, I. Alvarez-Martins, S.M. Gouveia, S.C. Jana, P. Duarte, T. Amado, P. Branco, M.C. Freitas, S.T. Silva, C. Antony, T.M. Bandeiras, and M. Bettencourt-Dias. 2012. BLD10/CEP135 Is a Microtubule-Associated Protein that Controls the Formation of the Flagellum Central Microtubule Pair. Developmental cell. 23:412-424.

Carvalho-Santos, Z., P. Machado, P. Branco, F. Tavares-Cadete, A. Rodrigues-Martins, J.B. Pereira-Leal, and M. Bettencourt-Dias. 2010. Stepwise evolution of the centriole- assembly pathway. Journal of cell science. 123:1414-1426.

120

Castresana, J. 2000. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Molecular biology and evolution. 17:540-552.

Cavalier-Smith, T. 1974. Basal body and flagellar development during the vegetative cell cycle and the sexual cycle of Chlamydomonas reinhardii. Journal of cell science. 16:529-556.

Cavalier-Smith, T. 1993. Kingdom protozoa and its 18 phyla. Microbiological reviews. 57:953-994.

Chakraborti, S., K. Natarajan, J. Curiel, C. Janke, and J. Liu. 2016. The emerging role of the tubulin code: From the tubulin molecule to neuronal function and disease. Cytoskeleton (Hoboken).

Chang, J.P., H. Mayahara, M. Yoloyama, A. Ubukata, and P.C. Moller. 1979. An ultrastructural study of morphogenesis of fibrogranular complex and centriole in ductuli efferentes of Chinese hamster. Tissue & cell. 11:401-412.

Chang, P., T.H. Giddings, Jr., M. Winey, and T. Stearns. 2003. Epsilon-tubulin is required for centriole duplication and microtubule organization. Nature cell biology. 5:71-76.

Chang, P., and T. Stearns. 2000. Delta-tubulin and epsilon-tubulin: two new human centrosomal tubulins reveal new aspects of centrosome structure and function. Nature cell biology. 2:30-35.

Cheeseman, I.M., C. Brew, M. Wolyniak, A. Desai, S. Anderson, N. Muster, J.R. Yates, T.C. Huffaker, D.G. Drubin, and G. Barnes. 2001. Implication of a novel multiprotein Dam1p complex in outer kinetochore function. The Journal of cell biology. 155:1137- 1145.

Cheeseman, I.M., and A. Desai. 2005. A combined approach for the localization and tandem affinity purification of protein complexes from metazoans. Science's STKE : signal transduction knowledge environment. 2005:pl1.

Cheeseman, I.M., S. Niessen, S. Anderson, F. Hyndman, J.R. Yates, 3rd, K. Oegema, and A. Desai. 2004. A conserved protein network controls assembly of the outer kinetochore and its ability to sustain tension. Genes & development. 18:2255-2268.

Chen, C.T., H. Hehnly, Q. Yu, D. Farkas, G. Zheng, S.D. Redick, H.F. Hung, R. Samtani, A. Jurczyk, S. Akbarian, C. Wise, A. Jackson, M. Bober, Y. Guo, C. Lo, and S. Doxsey. 2014. A unique set of centrosome proteins requires pericentrin for spindle-pole localization and spindle orientation. Current biology : CB. 24:2327-2334.

Chen, Z., V.B. Indjeian, M. McManus, L. Wang, and B.D. Dynlacht. 2002. CP110, a cell cycle-dependent CDK substrate, regulates centrosome duplication in human cells. Developmental cell. 3:339-350.

Cheng, J., N. Turkel, N. Hemati, M.T. Fuller, A.J. Hunt, and Y.M. Yamashita. 2008. Centrosome misorientation reduces stem cell division during ageing. Nature. 456:599-604.

Chevenet, F., C. Brun, A.L. Banuls, B. Jacq, and R. Christen. 2006. TreeDyn: towards dynamic graphics and annotations for analyses of trees. BMC bioinformatics. 7:439.

121

Chilvers, M.A., and C. O'Callaghan. 2000. Analysis of ciliary beat pattern and beat frequency using digital high speed imaging: comparison with the photomultiplier and photodiode methods. Thorax. 55:314-317.

Chretien, D., B. Buendia, S.D. Fuller, and E. Karsenti. 1997. Reconstruction of the centrosome cycle from cryoelectron micrographs. Journal of structural biology. 120:117-133.

Cole, E.S., K.R. Stuart, T.C. Marsh, K. Aufderheide, and W. Ringlien. 2002. Confocal fluorescence microscopy for Tetrahymena thermophila. Methods in cell biology. 70:337-359.

Culver, B.P., J.B. Meehl, T.H. Giddings, Jr., and M. Winey. 2009. The two SAS-6 homologs in Tetrahymena thermophila have distinct functions in basal body assembly. Molecular biology of the cell. 20:1865-1877.

Dahl, K.D., D.G. Sankaran, B.A. Bayless, M.E. Pinter, D.F. Galati, L.R. Heasley, T.H. Giddings, Jr., and C.G. Pearson. 2015. A Short CEP135 Splice Isoform Controls Centriole Duplication. Current biology : CB. 25:2591-2596.

Dahl, R., and L.A. Staehelin. 1989. High-pressure freezing for the preservation of biological structure: theory and practice. Journal of electron microscopy technique. 13:165-174.

Dammermann, A., P.S. Maddox, A. Desai, and K. Oegema. 2008. SAS-4 is recruited to a dynamic structure in newly forming centrioles that is stabilized by the gamma-tubulin- mediated addition of centriolar microtubules. The Journal of cell biology. 180:771- 785.

Delattre, M., S. Leidel, K. Wani, K. Baumer, J. Bamat, H. Schnabel, R. Feichtinger, R. Schnabel, and P. Gonczy. 2004. Centriolar SAS-5 is required for centrosome duplication in C. elegans. Nature cell biology. 6:656-664.

Delaval, B., A. Bright, N.D. Lawson, and S. Doxsey. 2011. The cilia protein IFT88 is required for spindle orientation in mitosis. Nature cell biology. 13:461-468.

Dereeper, A., V. Guignon, G. Blanc, S. Audic, S. Buffet, F. Chevenet, J.F. Dufayard, S. Guindon, V. Lefort, M. Lescot, J.M. Claverie, and O. Gascuel. 2008. Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic acids research. 36:W465- 469.

DiBella, L.M., and S.M. King. 2001. Dynein motors of the Chlamydomonas flagellum. International review of cytology. 210:227-268.

Dippell, R.V. 1967. How ciliary Basal bodies develop. Science. 158:527.

Dippell, R.V. 1968. The development of basal bodies in paramecium. Proceedings of the National Academy of Sciences of the United States of America. 61:461-468.

Dirksen, E.R. 1971. Centriole morphogenesis in developing ciliated epithelium of the mouse oviduct. The Journal of cell biology. 51:286-302.

Doxsey, S.J., P. Stein, L. Evans, P.D. Calarco, and M. Kirschner. 1994. Pericentrin, a highly conserved centrosome protein involved in microtubule organization. Cell. 76:639- 650.

122

Dupuis-Williams, P., A. Fleury-Aubusson, N.G. de Loubresse, H. Geoffroy, L. Vayssie, A. Galvani, A. Espigat, and J. Rossier. 2002. Functional role of epsilon-tubulin in the assembly of the centriolar microtubule scaffold. The Journal of cell biology. 158:1183-1193.

Dutcher, S.K., N.S. Morrissette, A.M. Preble, C. Rackley, and J. Stanga. 2002. Epsilon- tubulin is an essential component of the centriole. Molecular biology of the cell. 13:3859-3869.

Dutcher, S.K., and E.C. Trabuco. 1998. The UNI3 gene is required for assembly of basal bodies of Chlamydomonas and encodes delta-tubulin, a new member of the tubulin superfamily. Molecular biology of the cell. 9:1293-1308.

Edgar, R.C. 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic acids research. 32:1792-1797.

Eisen, J.A., R.S. Coyne, M. Wu, D. Wu, M. Thiagarajan, J.R. Wortman, J.H. Badger, Q. Ren, P. Amedeo, K.M. Jones, L.J. Tallon, A.L. Delcher, S.L. Salzberg, J.C. Silva, B.J. Haas, W.H. Majoros, M. Farzad, J.M. Carlton, R.K. Smith, Jr., J. Garg, R.E. Pearlman, K.M. Karrer, L. Sun, G. Manning, N.C. Elde, A.P. Turkewitz, D.J. Asai, D.E. Wilkes, Y. Wang, H. Cai, K. Collins, B.A. Stewart, S.R. Lee, K. Wilamowska, Z. Weinberg, W.L. Ruzzo, D. Wloga, J. Gaertig, J. Frankel, C.C. Tsao, M.A. Gorovsky, P.J. Keeling, R.F. Waller, N.J. Patron, J.M. Cherry, N.A. Stover, C.J. Krieger, C. del Toro, H.F. Ryder, S.C. Williamson, R.A. Barbeau, E.P. Hamilton, and E. Orias. 2006. Macronuclear genome sequence of the ciliate Tetrahymena thermophila, a model eukaryote. PLoS biology. 4:e286.

Fisch, C., and P. Dupuis-Williams. 2011. Ultrastructure of cilia and flagella - back to the future! Biology of the cell / under the auspices of the European Cell Biology Organization. 103:249-270.

Fouquet, J.P., Y. Prigent, and M.L. Kann. 1996. Comparative immunogold analysis of tubulin isoforms in the mouse sperm flagellum: unique distribution of glutamylated tubulin. Molecular reproduction and development. 43:358-365.

Fracek, S., and L. Margulis. 1979. Colchicine, nocodazole and trifluralin: different effects of microtubule polymerization inhibitors on the uptake and migration of endosymbiotic algae in Hydra viridis. Cytobios. 25:7-16.

Fromherz, S., T.H. Giddings, Jr., N. Gomez-Ospina, and S.K. Dutcher. 2004. Mutations in alpha-tubulin promote basal body maturation and flagellar assembly in the absence of delta-tubulin. Journal of cell science. 117:303-314.

Fujimura, M., and M. Okuno. 2006. Requirement of the fixed end for spontaneous beating in flagella. The Journal of experimental biology. 209:1336-1343.

Gaertig, J., L. Gu, B. Hai, and M.A. Gorovsky. 1994. High frequency vector-mediated transformation and gene replacement in Tetrahymena. Nucleic acids research. 22:5391-5398.

123

Galati, D.F., S. Bonney, Z. Kronenberg, C. Clarissa, M. Yandell, N.C. Elde, M. Jerka- Dziadosz, T.H. Giddings, J. Frankel, and C.G. Pearson. 2014. DisAp-dependent striated fiber elongation is required to organize ciliary arrays. The Journal of cell biology. 207:705-715.

Ganem, N.J., S.A. Godinho, and D. Pellman. 2009. A mechanism linking extra centrosomes to chromosomal instability. Nature. 460:278-282.

Garcia-Gonzalo, F.R., and J.F. Reiter. 2012. Scoring a backstage pass: Mechanisms of ciliogenesis and ciliary access. The Journal of cell biology. 197:697-709.

Gardner, L.C., E. O'Toole, C.A. Perrone, T. Giddings, and M.E. Porter. 1994. Components of a "dynein regulatory complex" are located at the junction between the radial spokes and the dynein arms in Chlamydomonas flagella. The Journal of cell biology. 127:1311-1325.

Garreau de Loubresse, N., F. Ruiz, J. Beisson, and C. Klotz. 2001. Role of delta-tubulin and the C-tubule in assembly of Paramecium basal bodies. BMC cell biology. 2:4.

Gibbons, I.R. 1963. Studies on the Protein Components of Cilia from Tetrahymena Pyriformis. Proceedings of the National Academy of Sciences of the United States of America. 50:1002-1010.

Gibbons, I.R. 1981. Cilia and flagella of eukaryotes. The Journal of cell biology. 91:107s- 124s.

Gibbons, I.R., and A.V. Grimstone. 1960. On flagellar structure in certain flagellates. The Journal of biophysical and biochemical cytology. 7:697-716.

Gibbons, I.R., and A.J. Rowe. 1965. Dynein: A Protein with Adenosine Triphosphatase Activity from Cilia. Science. 149:424-426.

Goetz, S.C., and K.V. Anderson. 2010. The primary cilium: a signalling centre during vertebrate development. Nature reviews. Genetics. 11:331-344.

Gogendeau, D., and R. Basto. 2010. Centrioles in flies: the exception to the rule? Seminars in cell & developmental biology. 21:163-173.

Gonczy, P. 2008. Mechanisms of asymmetric cell division: flies and worms pave the way. Nature reviews. Molecular cell biology. 9:355-366.

Goto, M., K. Ohki, and Y. Nozawa. 1982. Evidence for a correlation between swimming velocity and membrane fluidity of Tetrahymena cells. Biochimica et biophysica acta. 693:335-340.

Graser, S., Y.D. Stierhof, S.B. Lavoie, O.S. Gassner, S. Lamla, M. Le Clech, and E.A. Nigg. 2007. Cep164, a novel centriole appendage protein required for primary cilium formation. The Journal of cell biology. 179:321-330.

Greenstone, M.A., R.W. Jones, A. Dewar, B.G. Neville, and P.J. Cole. 1984. Hydrocephalus and primary ciliary dyskinesia. Archives of disease in childhood. 59:481-482.

Grill, S.W., J. Howard, E. Schaffer, E.H. Stelzer, and A.A. Hyman. 2003. The distribution of active force generators controls mitotic spindle position. Science. 301:518-521.

124

Grishchuk, E.L., M.I. Molodtsov, F.I. Ataullakhanov, and J.R. McIntosh. 2005. Force production by disassembling microtubules. Nature. 438:384-388.

Gromley, A., A. Jurczyk, J. Sillibourne, E. Halilovic, M. Mogensen, I. Groisman, M. Blomberg, and S. Doxsey. 2003. A novel human protein of the maternal centriole is required for the final stages of cytokinesis and entry into S phase. The Journal of cell biology. 161:535-545.

Guarguaglini, G., P.I. Duncan, Y.D. Stierhof, T. Holmstrom, S. Duensing, and E.A. Nigg. 2005. The forkhead-associated domain protein Cep170 interacts with Polo-like kinase 1 and serves as a marker for mature centrioles. Molecular biology of the cell. 16:1095-1107.

Gudi, R., C. Zou, J. Li, and Q. Gao. 2011. Centrobin-tubulin interaction is required for centriole elongation and stability. The Journal of cell biology. 193:711-725.

Guichard, P., D. Chretien, S. Marco, and A.M. Tassin. 2010. Procentriole assembly revealed by cryo-electron tomography. The EMBO journal. 29:1565-1572.

Guichard, P., A. Desfosses, A. Maheshwari, V. Hachet, C. Dietrich, A. Brune, T. Ishikawa, C. Sachse, and P. Gonczy. 2012. Cartwheel architecture of Trichonympha basal body. Science. 337:553.

Guichard, P., and P. Gonczy. 2016. Basal body structure in Trichonympha. Cilia. 5:9.

Guichard, P., V. Hachet, N. Majubu, A. Neves, D. Demurtas, N. Olieric, I. Fluckiger, A. Yamada, K. Kihara, Y. Nishida, S. Moriya, M.O. Steinmetz, Y. Hongoh, and P. Gonczy. 2013. Native architecture of the centriole proximal region reveals features underlying its 9-fold radial symmetry. Current biology : CB. 23:1620-1628.

Guindon, S., and O. Gascuel. 2003. A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Systematic biology. 52:696-704.

Habedanck, R., Y.D. Stierhof, C.J. Wilkinson, and E.A. Nigg. 2005. The Polo kinase Plk4 functions in centriole duplication. Nature cell biology. 7:1140-1146.

Hai, B., J. Gaertig, and M.A. Gorovsky. 2000. Knockout heterokaryons enable facile mutagenic analysis of essential genes in Tetrahymena. Methods in cell biology. 62:513-531.

Hata, H., Y. Yano, T. Mohri, H. Mohri, and T. Miki-Noumura. 1980. ATP-driven tubule extrusion from axonemes without outer dynein arms of sea-urchin sperm flagella. Journal of cell science. 41:331-340.

Hatch, E.M., A. Kulukian, A.J. Holland, D.W. Cleveland, and T. Stearns. 2010. Cep152 interacts with Plk4 and is required for centriole duplication. The Journal of cell biology. 191:721-729.

Heuser, T., M. Raytchev, J. Krell, M.E. Porter, and D. Nicastro. 2009. The dynein regulatory complex is the nexin link and a major regulatory node in cilia and flagella. The Journal of cell biology. 187:921-933.

125

Hilbert, M., A. Noga, D. Frey, V. Hamel, P. Guichard, S.H. Kraatz, M. Pfreundschuh, S. Hosner, I. Fluckiger, R. Jaussi, M.M. Wieser, K.M. Thieltges, X. Deupi, D.J. Muller, R.A. Kammerer, P. Gonczy, M. Hirono, and M.O. Steinmetz. 2016. SAS-6 engineering reveals interdependence between cartwheel and microtubules in determining centriole architecture. Nature cell biology. 18:393-403.

Hildebrandt, F., T. Benzing, and N. Katsanis. 2011. Ciliopathies. The New England journal of medicine. 364:1533-1543.

Hill, D.B., V. Swaminathan, A. Estes, J. Cribb, E.T. O'Brien, C.W. Davis, and R. Superfine. 2010. Force generation and dynamics of individual cilia under external loading. Biophysical journal. 98:57-66.

Hinchcliffe, E.H., F.J. Miller, M. Cham, A. Khodjakov, and G. Sluder. 2001. Requirement of a centrosomal activity for cell cycle progression through G1 into S phase. Science. 291:1547-1550.

Hiraki, M., Y. Nakazawa, R. Kamiya, and M. Hirono. 2007. Bld10p constitutes the cartwheel- spoke tip and stabilizes the 9-fold symmetry of the centriole. Current biology : CB. 17:1778-1783.

Hodges, M.E., N. Scheumann, B. Wickstead, J.A. Langdale, and K. Gull. 2010. Reconstructing the evolutionary history of the centriole from protein components. Journal of cell science. 123:1407-1413.

Holland, A.J., W. Lan, and D.W. Cleveland. 2010. Centriole duplication: A lesson in self- control. Cell Cycle. 9:2731-2736.

Holmes, J.A., and S.K. Dutcher. 1989. Cellular asymmetry in Chlamydomonas reinhardtii. Journal of cell science. 94 ( Pt 2):273-285.

Holzbaur, E.L., and R.B. Vallee. 1994. DYNEINS: molecular structure and cellular function. Annual review of cell biology. 10:339-372.

Hoppeler-Lebel, A., C. Celati, G. Bellett, M.M. Mogensen, L. Klein-Hitpass, M. Bornens, and A.M. Tassin. 2007. Centrosomal CAP350 protein stabilises microtubules associated with the Golgi complex. Journal of cell science. 120:3299-3308.

Howes, S.C., G.M. Alushin, T. Shida, M.V. Nachury, and E. Nogales. 2014. Effects of tubulin acetylation and tubulin acetyltransferase binding on microtubule structure. Molecular biology of the cell. 25:257-266.

Hozumi, A., Y. Satouh, Y. Makino, T. Toda, H. Ide, K. Ogawa, S.M. King, and K. Inaba. 2006. Molecular characterization of Ciona sperm outer arm dynein reveals multiple components related to outer arm docking complex protein 2. Cell motility and the cytoskeleton. 63:591-603.

Hsu, W.B., L.Y. Hung, C.J. Tang, C.L. Su, Y. Chang, and T.K. Tang. 2008. Functional characterization of the microtubule-binding and -destabilizing domains of CPAP and d-SAS-4. Experimental cell research. 314:2591-2602.

126

Huitorel, P., D. White, J.P. Fouquet, M.L. Kann, J. Cosson, and C. Gagnon. 2002. Differential distribution of glutamylated tubulin isoforms along the sea urchin sperm axoneme. Molecular reproduction and development. 62:139-148.

Hung, H.F., H. Hehnly, and S. Doxsey. 2016. The Mother Centriole Appendage Protein Cenexin Modulates Lumen Formation through Spindle Orientation. Current biology : CB. 26:793-801.

Hussain, M.S., S.M. Baig, S. Neumann, G. Nurnberg, M. Farooq, I. Ahmad, T. Alef, H.C. Hennies, M. Technau, J. Altmuller, P. Frommolt, H. Thiele, A.A. Noegel, and P. Nurnberg. 2012. A Truncating Mutation of CEP135 Causes Primary Microcephaly and Disturbed Centrosomal Function. American journal of human genetics. 90:871- 878.

Hyman, A.A. 1989. Centrosome movement in the early divisions of Caenorhabditis elegans: a cortical site determining centrosome position. The Journal of cell biology. 109:1185-1193.

Ibanez-Tallon, I., S. Gorokhova, and N. Heintz. 2002. Loss of function of axonemal dynein Mdnah5 causes primary ciliary dyskinesia and hydrocephalus. Human molecular genetics. 11:715-721.

Iftode, F., and A. Fleury-Aubusson. 2003. Structural inheritance in Paramecium: ultrastructural evidence for basal body and associated rootlets polarity transmission through binary fission. Biology of the cell / under the auspices of the European Cell Biology Organization. 95:39-51.

Ikegami, K., S. Sato, K. Nakamura, L.E. Ostrowski, and M. Setou. 2010. Tubulin polyglutamylation is essential for airway ciliary function through the regulation of beating asymmetry. Proceedings of the National Academy of Sciences of the United States of America. 107:10490-10495.

Inaba, K. 2007. Molecular basis of sperm flagellar axonemes: structural and evolutionary aspects. Annals of the New York Academy of Sciences. 1101:506-526.

Inclan, Y.F., and E. Nogales. 2001. Structural models for the self-assembly and microtubule interactions of gamma-, delta- and epsilon-tubulin. Journal of cell science. 114:413- 422.

Ishijima, S. 2012. Mechanical constraint converts planar waves into helices on tunicate and sea urchin sperm flagella. Cell structure and function. 37:13-19.

Ishijima, S., M.S. Hamaguchi, M. Naruse, S.A. Ishijima, and Y. Hamaguchi. 1992. Rotational movement of a spermatozoon around its long axis. The Journal of experimental biology. 163:15-31.

Ishijima, S., and Y. Hamaguchi. 1993. Calcium ion regulation of chirality of beating flagellum of reactivated sea urchin spermatozoa. Biophysical journal. 65:1445-1448.

Ishijima, S., and Y. Hiramoto. 1994. Flexural rigidity of echinoderm sperm flagella. Cell structure and function. 19:349-362.

127

Ishijima, S., M. Kubo-Irie, H. Mohri, and Y. Hamaguchi. 1996. Calcium-dependent bidirectional power stroke of the dynein arms in sea urchin sperm axonemes. Journal of cell science. 109 ( Pt 12):2833-2842.

Izquierdo, D., W.J. Wang, K. Uryu, and M.F. Tsou. 2014. Stabilization of cartwheel-less centrioles for duplication requires CEP295-mediated centriole-to-centrosome conversion. Cell reports. 8:957-965.

Jackman, M., C. Lindon, E.A. Nigg, and J. Pines. 2003. Active cyclin B1-Cdk1 first appears on centrosomes in prophase. Nature cell biology. 5:143-148.

Janke, C., K. Rogowski, D. Wloga, C. Regnard, A.V. Kajava, J.M. Strub, N. Temurak, J. van Dijk, D. Boucher, A. van Dorsselaer, S. Suryavanshi, J. Gaertig, and B. Edde. 2005. Tubulin polyglutamylase enzymes are members of the TTL domain protein family. Science. 308:1758-1762.

Jeong, Y., J. Lee, K. Kim, J.C. Yoo, and K. Rhee. 2007. Characterization of NIP2/centrobin, a novel substrate of Nek2, and its potential role in microtubule stabilization. Journal of cell science. 120:2106-2116.

Jerka-Dziadosz, M. 1981. Cytoskeleton-related structures in tetrahymena thermophila: microfilaments at the apical and division-furrow rings. Journal of cell science. 51:241- 253.

Jerka-Dziadosz, M., D. Gogendeau, C. Klotz, J. Cohen, J. Beisson, and F. Koll. 2010. Basal body duplication in Paramecium: the key role of Bld10 in assembly and stability of the cartwheel. Cytoskeleton (Hoboken). 67:161-171.

Jerka-Dziadosz, M., L.M. Jenkins, E.M. Nelsen, N.E. Williams, R. Jaeckel-Williams, and J. Frankel. 1995. Cellular polarity in ciliates: persistence of global polarity in a disorganized mutant of Tetrahymena thermophila that disrupts cytoskeletal organization. Developmental biology. 169:644-661.

Jerka-Dziadosz, M., F. Koll, D. Wloga, D. Gogendeau, N. Garreau de Loubresse, F. Ruiz, S. Fabczak, and J. Beisson. 2013. A Centrin3-dependent, transient, appendage of the mother basal body guides the positioning of the daughter basal body in Paramecium. Protist. 164:352-368.

Jung, I., T.R. Powers, and J.M. Valles, Jr. 2014. Evidence for two extremes of ciliary motor response in a single swimming microorganism. Biophysical journal. 106:106-113.

Kagami, O., S. Takada, and R. Kamiya. 1990. Microtubule translocation caused by three subspecies of inner-arm dynein from Chlamydomonas flagella. FEBS letters. 264:179-182.

Kalnins, V.I., and K.R. Porter. 1969. Centriole replication during ciliogenesis in the chick tracheal epithelium. Z Zellforsch Mikrosk Anat. 100:1-30.

Kanous, K.S., C. Casey, and C.B. Lindemann. 1993. Inhibition of microtubule sliding by Ni2+ and Cd2+: evidence for a differential response of certain microtubule pairs within the bovine sperm axoneme. Cell motility and the cytoskeleton. 26:66-76.

128

Khodjakov, A., R.W. Cole, B.R. Oakley, and C.L. Rieder. 2000. Centrosome-independent mitotic spindle formation in vertebrates. Current biology : CB. 10:59-67.

Khodjakov, A., and C.L. Rieder. 2001. Centrosomes enhance the fidelity of cytokinesis in vertebrates and are required for cell cycle progression. The Journal of cell biology. 153:237-242.

Kilburn, C.L., C.G. Pearson, E.P. Romijn, J.B. Meehl, T.H. Giddings, Jr., B.P. Culver, J.R. Yates, 3rd, and M. Winey. 2007. New Tetrahymena basal body protein components identify basal body domain structure. The Journal of cell biology. 178:905-912.

Kim, H. 1991. Depletion of acetylated alpha-tubulin during microtubule purification from bovine brain gray and white matter regions. Journal of neuroscience research. 30:172-182.

Kim, T.S., J.E. Park, A. Shukla, S. Choi, R.N. Murugan, J.H. Lee, M. Ahn, K. Rhee, J.K. Bang, B.Y. Kim, J. Loncarek, R.L. Erikson, and K.S. Lee. 2013. Hierarchical recruitment of Plk4 and regulation of centriole biogenesis by two centrosomal scaffolds, Cep192 and Cep152. Proceedings of the National Academy of Sciences of the United States of America. 110:E4849-4857.

Kinukawa, M., S. Oda, Y. Shirakura, M. Okabe, J. Ohmuro, S.A. Baba, M. Nagata, and F. Aoki. 2006. Roles of cAMP in regulating microtubule sliding and flagellar bending in demembranated hamster spermatozoa. FEBS letters. 580:1515-1520.

Kitagawa, D., I. Vakonakis, N. Olieric, M. Hilbert, D. Keller, V. Olieric, M. Bortfeld, M.C. Erat, I. Fluckiger, P. Gonczy, and M.O. Steinmetz. 2011. Structural basis of the 9-fold symmetry of centrioles. Cell. 144:364-375.

Kleylein-Sohn, J., J. Westendorf, M. Le Clech, R. Habedanck, Y.D. Stierhof, and E.A. Nigg. 2007. Plk4-induced centriole biogenesis in human cells. Developmental cell. 13:190- 202.

Klos Dehring, D.A., E.K. Vladar, M.E. Werner, J.W. Mitchell, P. Hwang, and B.J. Mitchell. 2013. Deuterosome-mediated centriole biogenesis. Developmental cell. 27:103-112.

Kobayashi, Y., and H. Mohri. 1977. Microheterogeneity of alpha and beta subunit of tubulin from microtubules of starfish (Asterias amurensis) sperm flagella. Journal of molecular biology. 116:613-617.

Kochanski, R.S., and G.G. Borisy. 1990. Mode of centriole duplication and distribution. The Journal of cell biology. 110:1599-1605.

Kohlmaier, G., J. Loncarek, X. Meng, B.F. McEwen, M.M. Mogensen, A. Spektor, B.D. Dynlacht, A. Khodjakov, and P. Gonczy. 2009. Overly long centrioles and defective cell division upon excess of the SAS-4-related protein CPAP. Current biology : CB. 19:1012-1018.

Kong, D., V. Farmer, A. Shukla, J. James, R. Gruskin, S. Kiriyama, and J. Loncarek. 2014. Centriole maturation requires regulated Plk1 activity during two consecutive cell cycles. The Journal of cell biology. 206:855-865.

129

Kozlowski, C., M. Srayko, and F. Nedelec. 2007. Cortical microtubule contacts position the spindle in C. elegans embryos. Cell. 129:499-510.

Kratz, A.S., F. Barenz, K.T. Richter, and I. Hoffmann. 2015. Plk4-dependent phosphorylation of STIL is required for centriole duplication. Biol Open. 4:370-377.

Kubai, D.F. 1975. The evolution of the mitotic spindle. International review of cytology. 43:167-227.

Kubo, T., T. Yagi, and R. Kamiya. 2012. Tubulin polyglutamylation regulates flagellar motility by controlling a specific inner-arm dynein that interacts with the dynein regulatory complex. Cytoskeleton (Hoboken). 69:1059-1068.

Kubo, T., H.A. Yanagisawa, T. Yagi, M. Hirono, and R. Kamiya. 2010. Tubulin polyglutamylation regulates axonemal motility by modulating activities of inner-arm dyneins. Current biology : CB. 20:441-445.

Kuijpers, M., and C.C. Hoogenraad. 2011. Centrosomes, microtubules and neuronal development. Molecular and cellular neurosciences. 48:349-358.

Kunimoto, K., Y. Yamazaki, T. Nishida, K. Shinohara, H. Ishikawa, T. Hasegawa, T. Okanoue, H. Hamada, T. Noda, A. Tamura, and S. Tsukita. 2012. Coordinated ciliary beating requires Odf2-mediated polarization of basal bodies via basal feet. Cell. 148:189-200.

La Terra, S., C.N. English, P. Hergert, B.F. McEwen, G. Sluder, and A. Khodjakov. 2005. The de novo centriole assembly pathway in HeLa cells: cell cycle progression and centriole assembly/maturation. The Journal of cell biology. 168:713-722.

Lacroix, B., J. van Dijk, N.D. Gold, J. Guizetti, G. Aldrian-Herrada, K. Rogowski, D.W. Gerlich, and C. Janke. 2010. Tubulin polyglutamylation stimulates spastin-mediated microtubule severing. The Journal of cell biology. 189:945-954.

Lange, B.M., and K. Gull. 1995. A molecular marker for centriole maturation in the mammalian cell cycle. The Journal of cell biology. 130:919-927.

Larsen, J., and P. Satir. 1991. Analysis of Ni(2+)-induced arrest of Paramecium axonemes. Journal of cell science. 99 ( Pt 1):33-40.

Lawo, S., M. Hasegan, G.D. Gupta, and L. Pelletier. 2012. Subdiffraction imaging of centrosomes reveals higher-order organizational features of pericentriolar material. Nature cell biology. 14:1148-1158.

Le Clech, M. 2008. Role of CAP350 in centriolar tubule stability and centriole assembly. PloS one. 3:e3855.

Lechtreck, K.F., and M. Melkonian. 1991. Striated microtubule-associated fibers: identification of assemblin, a novel 34-kD protein that forms paracrystals of 2-nm filaments in vitro. The Journal of cell biology. 115:705-716.

Lechtreck, K.F., and M. Melkonian. 1998. SF-assemblin, striated fibers, and segmented coiled coil proteins. Cell motility and the cytoskeleton. 41:289-296.

130

Lee, K., and K. Rhee. 2011. PLK1 phosphorylation of pericentrin initiates centrosome maturation at the onset of mitosis. The Journal of cell biology. 195:1093-1101.

Li, S., J.J. Fernandez, W.F. Marshall, and D.A. Agard. 2012. Three-dimensional structure of basal body triplet revealed by electron cryo-tomography. The EMBO journal. 31:552- 562.

Lin, Y.C., C.W. Chang, W.B. Hsu, C.J. Tang, Y.N. Lin, E.J. Chou, C.T. Wu, and T.K. Tang. 2013. Human microcephaly protein CEP135 binds to hSAS-6 and CPAP, and is required for centriole assembly. The EMBO journal. 32:1141-1154.

Lindemann, C.B., and K.S. Kanous. 1997. A model for flagellar motility. International review of cytology. 173:1-72.

Lindemann, C.B., A. Orlando, and K.S. Kanous. 1992. The flagellar beat of rat sperm is organized by the interaction of two functionally distinct populations of dynein bridges with a stable central axonemal partition. Journal of cell science. 102 ( Pt 2):249-260.

Loktev, A.V., Q. Zhang, J.S. Beck, C.C. Searby, T.E. Scheetz, J.F. Bazan, D.C. Slusarski, V.C. Sheffield, P.K. Jackson, and M.V. Nachury. 2008. A BBSome subunit links ciliogenesis, microtubule stability, and acetylation. Developmental cell. 15:854-865.

Lopes, C.A., S.C. Jana, I. Cunha-Ferreira, S. Zitouni, I. Bento, P. Duarte, S. Gilberto, F. Freixo, A. Guerrero, M. Francia, M. Lince-Faria, J. Carneiro, and M. Bettencourt- Dias. 2015. PLK4 trans-Autoactivation Controls Centriole Biogenesis in Space. Developmental cell. 35:222-235.

Lorch, D.P., C.B. Lindemann, and A.J. Hunt. 2008. The motor activity of mammalian axonemal dynein studied in situ on doublet microtubules. Cell motility and the cytoskeleton. 65:487-494.

Lyons, R.A., E. Saridogan, and O. Djahanbakhch. 2006. The reproductive significance of human Fallopian tube cilia. Human reproduction update. 12:363-372.

Machemer, H. 1972. Ciliary activity and the origin of metachrony in Paramecium: effects of increased viscosity. The Journal of experimental biology. 57:239-259.

Magiera, M.M., and C. Janke. 2014. Post-translational modifications of tubulin. Current biology : CB. 24:R351-354.

Manandhar, G., C. Simerly, J.L. Salisbury, and G. Schatten. 1999. Centriole and centrin degeneration during mouse spermiogenesis. Cell motility and the cytoskeleton. 43:137-144.

Maniotis, A., and M. Schliwa. 1991. Microsurgical removal of centrosomes blocks cell reproduction and centriole generation in BSC-1 cells. Cell. 67:495-504.

Marshall, W.F. 2009. Centriole evolution. Current opinion in cell biology. 21:14-19.

Marszalek, J.R., X. Liu, E.A. Roberts, D. Chui, J.D. Marth, D.S. Williams, and L.S. Goldstein. 2000. Genetic evidence for selective transport of opsin and arrestin by kinesin-II in mammalian photoreceptors. Cell. 102:175-187.

131

Maruta, H., K. Greer, and J.L. Rosenbaum. 1986. The acetylation of alpha-tubulin and its relationship to the assembly and disassembly of microtubules. The Journal of cell biology. 103:571-579.

Matsumoto, Y., and J.L. Maller. 2004. A centrosomal localization signal in cyclin E required for Cdk2-independent S phase entry. Science. 306:885-888.

Matsuura, K., P.A. Lefebvre, R. Kamiya, and M. Hirono. 2004. Bld10p, a novel protein essential for basal body assembly in Chlamydomonas: localization to the cartwheel, the first ninefold symmetrical structure appearing during assembly. The Journal of cell biology. 165:663-671.

Matsuyama, A., T. Shimazu, Y. Sumida, A. Saito, Y. Yoshimatsu, D. Seigneurin-Berny, H. Osada, Y. Komatsu, N. Nishino, S. Khochbin, S. Horinouchi, and M. Yoshida. 2002. In vivo destabilization of dynamic microtubules by HDAC6-mediated deacetylation. The EMBO journal. 21:6820-6831.

Meehl, J.B. 2016. Tetrahymena Poc1 ensures proper intertriplet microtubule linkages to maintain basal body integrity. Molecular biology of the cell. Jun 1.

Meehl, J.B., T.H. Giddings, Jr., and M. Winey. 2009. High pressure freezing, electron microscopy, and immuno-electron microscopy of Tetrahymena thermophila basal bodies. Methods Mol Biol. 586:227-241.

Mennella, V., D.A. Agard, B. Huang, and L. Pelletier. 2014. Amorphous no more: subdiffraction view of the pericentriolar material architecture. Trends in cell biology. 24:188-197.

Miao, W., J. Xiong, J. Bowen, W. Wang, Y. Liu, O. Braguinets, J. Grigull, R.E. Pearlman, E. Orias, and M.A. Gorovsky. 2009. Microarray analyses of gene expression during the Tetrahymena thermophila life cycle. PloS one. 4:e4429.

Mikolajka, A., X. Yan, G.M. Popowicz, P. Smialowski, E.A. Nigg, and T.A. Holak. 2006. Structure of the N-terminal domain of the FOP (FGFR1OP) protein and implications for its dimerization and centrosomal localization. Journal of molecular biology. 359:863-875.

Mikule, K., B. Delaval, P. Kaldis, A. Jurcyzk, P. Hergert, and S. Doxsey. 2007. Loss of centrosome integrity induces p38-p53-p21-dependent G1-S arrest. Nature cell biology. 9:160-170.

Mitchison, T., and M. Kirschner. 1984. Dynamic instability of microtubule growth. Nature. 312:237-242.

Mogensen, M.M., A. Malik, M. Piel, V. Bouckson-Castaing, and M. Bornens. 2000. Microtubule minus-end anchorage at centrosomal and non-centrosomal sites: the role of ninein. Journal of cell science. 113 ( Pt 17):3013-3023.

Mohri, H., K. Inaba, S. Ishijima, and S.A. Baba. 2012. Tubulin-dynein system in flagellar and ciliary movement. Proceedings of the Japan Academy. Series B, Physical and biological sciences. 88:397-415.

132

Moser, J.J., M.J. Fritzler, Y. Ou, and J.B. Rattner. 2010. The PCM-basal body/primary cilium coalition. Seminars in cell & developmental biology. 21:148-155.

Moss, A.G., W.S. Sale, L.A. Fox, and G.B. Witman. 1992. The alpha subunit of sea urchin sperm outer arm dynein mediates structural and rigor binding to microtubules. The Journal of cell biology. 118:1189-1200.

Mottier-Pavie, V., and T.L. Megraw. 2009. Drosophila bld10 is a centriolar protein that regulates centriole, basal body, and motile cilium assembly. Molecular biology of the cell. 20:2605-2614.

Moudjou, M., N. Bordes, M. Paintrand, and M. Bornens. 1996. gamma-Tubulin in mammalian cells: the centrosomal and the cytosolic forms. Journal of cell science. 109 ( Pt 4):875-887.

Moyer, T.C., K.M. Clutario, B.G. Lambrus, V. Daggubati, and A.J. Holland. 2015. Binding of STIL to Plk4 activates kinase activity to promote centriole assembly. The Journal of cell biology. 209:863-878.

Nachury, M.V., A.V. Loktev, Q. Zhang, C.J. Westlake, J. Peranen, A. Merdes, D.C. Slusarski, R.H. Scheller, J.F. Bazan, V.C. Sheffield, and P.K. Jackson. 2007. A core complex of BBS proteins cooperates with the GTPase Rab8 to promote ciliary membrane biogenesis. Cell. 129:1201-1213.

Nakagawa, Y., Y. Yamane, T. Okanoue, and S. Tsukita. 2001. Outer dense fiber 2 is a widespread centrosome scaffold component preferentially associated with mother centrioles: its identification from isolated centrosomes. Molecular biology of the cell. 12:1687-1697.

Nakano, I., T. Kobayashi, M. Yoshimura, and C. Shingyoji. 2003. Central-pair-linked regulation of microtubule sliding by calcium in flagellar axonemes. Journal of cell science. 116:1627-1636.

Nakazawa, Y., M. Hiraki, R. Kamiya, and M. Hirono. 2007. SAS-6 is a cartwheel protein that establishes the 9-fold symmetry of the centriole. Current biology : CB. 17:2169-2174.

Nanney, D.L., and M. Chow. 1974. Basal body homeostasis in Tetrahymena. Am. Nat.:125- 139.

Nauli, S.M., X. Jin, and B.P. Hierck. 2011. The mechanosensory role of primary cilia in vascular hypertension. International journal of vascular medicine. 2011:376281.

Ng, S.F., and J. Frankel. 1977. 180 degrees rotation of ciliary rows and its morphogenetic implications in Tetrahymena pyriformis. Proceedings of the National Academy of Sciences of the United States of America. 74:1115-1119.

Nguyen-Ngoc, T., K. Afshar, and P. Gonczy. 2007. Coupling of cortical dynein and G alpha proteins mediates spindle positioning in Caenorhabditis elegans. Nature cell biology. 9:1294-1302.

Nguyen, A.M., and C.R. Jacobs. 2013. Emerging role of primary cilia as mechanosensors in osteocytes. Bone. 54:196-204.

133

Nigg, E.A. 2007. Centrosome duplication: of rules and licenses. Trends in cell biology. 17:215-221.

Nigg, E.A., and J.W. Raff. 2009. Centrioles, centrosomes, and cilia in health and disease. Cell. 139:663-678.

Nigg, E.A., and T. Stearns. 2011. The centrosome cycle: Centriole biogenesis, duplication and inherent asymmetries. Nature cell biology. 13:1154-1160.

Nogales, E., S.G. Wolf, and K.H. Downing. 1998. Structure of the alpha beta tubulin dimer by electron crystallography. Nature. 391:199-203.

O'Toole, E.T., and S.K. Dutcher. 2014. Site-specific basal body duplication in Chlamydomonas. Cytoskeleton (Hoboken). 71:108-118.

O'Toole, E.T., T.H. Giddings, J.R. McIntosh, and S.K. Dutcher. 2003. Three-dimensional organization of basal bodies from wild-type and delta-tubulin deletion strains of Chlamydomonas reinhardtii. Molecular biology of the cell. 14:2999-3012.

Ogawa, K., T. Mohri, and H. Mohri. 1977. Identification of dynein as the outer arms of sea urchin sperm axonemes. Proceedings of the National Academy of Sciences of the United States of America. 74:5006-5010.

Ogawa, K., and H. Mori. 1975. Preparation of antiserum against a tryptic fragment (fragment A) of dynein and an immunological approach to the subunit composition of dynein. The Journal of biological chemistry. 250:6476-6483.

Ogawa, K., H. Takai, A. Ogiwara, E. Yokota, T. Shimizu, K. Inaba, and H. Mohri. 1996. Is outer arm dynein intermediate chain 1 multifunctional? Molecular biology of the cell. 7:1895-1907.

Palevitz, B.A. 1993. Morphological Plasticity of the Mitotic Apparatus in Plants and Its Developmental Consequences. The Plant cell. 5:1001-1009.

Pathak, N., C.A. Austin, and I.A. Drummond. 2011. Tubulin tyrosine ligase-like genes ttll3 and ttll6 maintain zebrafish cilia structure and motility. The Journal of biological chemistry. 286:11685-11695.

Pearson, C.G. 2014. Choosing sides--asymmetric centriole and basal body assembly. Journal of cell science. 127:2803-2810.

Pearson, C.G., B.P. Culver, and M. Winey. 2007. Centrioles want to move out and make cilia. Developmental cell. 13:319-321.

Pearson, C.G., T.H. Giddings, Jr., and M. Winey. 2009a. Basal body components exhibit differential protein dynamics during nascent basal body assembly. Molecular biology of the cell. 20:904-914.

Pearson, C.G., D.P. Osborn, T.H. Giddings, Jr., P.L. Beales, and M. Winey. 2009b. Basal body stability and ciliogenesis requires the conserved component Poc1. The Journal of cell biology. 187:905-920.

Pearson, C.G., and M. Winey. 2009. Basal body assembly in ciliates: the power of numbers. Traffic. 10:461-471.

134

Pelletier, L., E. O'Toole, A. Schwager, A.A. Hyman, and T. Muller-Reichert. 2006. Centriole assembly in Caenorhabditis elegans. Nature. 444:619-623.

Perlman, B.S. 1973. Basal body addition in ciliary rows of Tetrahymena pyriformis. The Journal of experimental zoology. 184:365-368.

Piperno, G., K. Mead, and W. Shestak. 1992. The inner dynein arms I2 interact with a "dynein regulatory complex" in Chlamydomonas flagella. The Journal of cell biology. 118:1455-1463.

Raynaud-Messina, B., L. Mazzolini, A. Moisand, A.M. Cirinesi, and M. Wright. 2004. Elongation of centriolar microtubule triplets contributes to the formation of the mitotic spindle in gamma-tubulin-depleted cells. Journal of cell science. 117:5497-5507.

Reed, N.A., D. Cai, T.L. Blasius, G.T. Jih, E. Meyhofer, J. Gaertig, and K.J. Verhey. 2006. Microtubule acetylation promotes kinesin-1 binding and transport. Current biology : CB. 16:2166-2172.

Riedel-Kruse, I.H., A. Hilfinger, J. Howard, and F. Julicher. 2007. How molecular motors shape the flagellar beat. HFSP journal. 1:192-208.

Roque, H., A. Wainman, J. Richens, K. Kozyrska, A. Franz, and J.W. Raff. 2012. Drosophila Cep135/Bld10 maintains proper centriole structure but is dispensable for cartwheel formation. Journal of cell science. 125:5881-5886.

Ross, I., C. Clarissa, T.H. Giddings, Jr., and M. Winey. 2013. epsilon-tubulin is essential in Tetrahymena thermophila for the assembly and stability of basal bodies. Journal of cell science. 126:3441-3451.

Roth, K.E., C.L. Rieder, and S.S. Bowser. 1988. Flexible-substratum technique for viewing cells from the side: some in vivo properties of primary (9+0) cilia in cultured kidney epithelia. Journal of cell science. 89 ( Pt 4):457-466.

Saeki, H., S. Kondo, T. Morita, I. Sasagawa, G. Ishizuka, and Y. Koizumi. 1984. Immotile cilia syndrome associated with polycystic kidney. The Journal of urology. 132:1165- 1166.

Sale, W.S. 1986. The axonemal axis and Ca2+-induced asymmetry of active microtubule sliding in sea urchin sperm tails. The Journal of cell biology. 102:2042-2052.

Sale, W.S., and P. Satir. 1977. Direction of active sliding of microtubules in Tetrahymena cilia. Proceedings of the National Academy of Sciences of the United States of America. 74:2045-2049.

Satir, P. 1965. STUDIES ON CILIA : II. Examination of the Distal Region of the Ciliary Shaft and the Role of the Filaments in Motility. The Journal of cell biology. 26:805-834.

Satir, P. 1968. Studies on cilia. 3. Further studies on the cilium tip and a "sliding filament" model of ciliary motility. The Journal of cell biology. 39:77-94.

Sawamoto, K., H. Wichterle, O. Gonzalez-Perez, J.A. Cholfin, M. Yamada, N. Spassky, N.S. Murcia, J.M. Garcia-Verdugo, O. Marin, J.L. Rubenstein, M. Tessier-Lavigne, H. Okano, and A. Alvarez-Buylla. 2006. New neurons follow the flow of cerebrospinal fluid in the adult brain. Science. 311:629-632.

135

Schmidt, T.I., J. Kleylein-Sohn, J. Westendorf, M. Le Clech, S.B. Lavoie, Y.D. Stierhof, and E.A. Nigg. 2009. Control of centriole length by CPAP and CP110. Current biology : CB. 19:1005-1011.

Schouteden, C., D. Serwas, M. Palfy, and A. Dammermann. 2015. The ciliary transition zone functions in cell adhesion but is dispensable for axoneme assembly in C. elegans. The Journal of cell biology. 210:35-44.

Shang, Y., B. Li, and M.A. Gorovsky. 2002. Tetrahymena thermophila contains a conventional gamma-tubulin that is differentially required for the maintenance of different microtubule-organizing centers. The Journal of cell biology. 158:1195-1206.

Shang, Y., C.C. Tsao, and M.A. Gorovsky. 2005. Mutational analyses reveal a novel function of the nucleotide-binding domain of gamma-tubulin in the regulation of basal body biogenesis. The Journal of cell biology. 171:1035-1044.

Sharma, N., J. Bryant, D. Wloga, R. Donaldson, R.C. Davis, M. Jerka-Dziadosz, and J. Gaertig. 2007. Katanin regulates dynamics of microtubules and biogenesis of motile cilia. The Journal of cell biology. 178:1065-1079.

Shingyoji, C., A. Murakami, and K. Takahashi. 1977. Local reactivation of Triton-extracted flagella by iontophoretic application of ATP. Nature. 265:269-270.

Shishikura, F., and K. Sekiguchi. 1979. Comparative studies on hemocytes and coagulogens of the Asian and the American horseshoe crabs. Progress in clinical and biological research. 29:185-201.

Silflow, C.D., and P.A. Lefebvre. 2001. Assembly and motility of eukaryotic cilia and flagella. Lessons from Chlamydomonas reinhardtii. Plant physiology. 127:1500-1507.

Silkworth, W.T., I.K. Nardi, L.M. Scholl, and D. Cimini. 2009. Multipolar spindle pole coalescence is a major source of kinetochore mis-attachment and chromosome mis- segregation in cancer cells. PloS one. 4:e6564.

Sir, J.H., M. Putz, O. Daly, C.G. Morrison, M. Dunning, J.V. Kilmartin, and F. Gergely. 2013. Loss of centrioles causes chromosomal instability in vertebrate somatic cells. The Journal of cell biology. 203:747-756.

Smith, E.F., and P. Yang. 2004. The radial spokes and central apparatus: mechano- chemical transducers that regulate flagellar motility. Cell motility and the cytoskeleton. 57:8-17.

Sonnen, K.F., A.M. Gabryjonczyk, E. Anselm, Y.D. Stierhof, and E.A. Nigg. 2013. Human Cep192 and Cep152 cooperate in Plk4 recruitment and centriole duplication. Journal of cell science. 126:3223-3233.

Sorokin, S.P. 1968a. Centriole formation and ciliogenesis. Aspen Emphysema Conference. 11:213-216.

Sorokin, S.P. 1968b. Reconstructions of centriole formation and ciliogenesis in mammalian lungs. Journal of cell science. 3:207-230.

Spektor, A., W.Y. Tsang, D. Khoo, and B.D. Dynlacht. 2007. Cep97 and CP110 suppress a cilia assembly program. Cell. 130:678-690.

136

Sperling, L., G. Keryer, F. Ruiz, and J. Beisson. 1991. Cortical morphogenesis in Paramecium: a transcellular wave of protein phosphorylation involved in ciliary rootlet disassembly. Developmental biology. 148:205-218.

Spoon, D.M., C.O. Feise, II, and R.S. Youn. 1977. Poly(ethylene oxide), a new slowing agent for protozoa. The Journal of protozoology. 24:471-474.

Steinman, R.M. 1968. An electron microscopic study of ciliogenesis in developing epidermis and trachea in the embryo of Xenopus laevis. The American journal of anatomy. 122:19-55.

Stemm-Wolf, A.J., J.B. Meehl, and M. Winey. 2013. Sfr13, a member of a large family of asymmetrically localized Sfi1-repeat proteins, is important for basal body separation and stability in Tetrahymena thermophila. Journal of cell science. 126:1659-1671.

Stemm-Wolf, A.J., G. Morgan, T.H. Giddings, Jr., E.A. White, R. Marchione, H.B. McDonald, and M. Winey. 2005. Basal body duplication and maintenance require one member of the Tetrahymena thermophila centrin gene family. Molecular biology of the cell. 16:3606-3619.

Stevens, N.R., J. Dobbelaere, K. Brunk, A. Franz, and J.W. Raff. 2010. Drosophila Ana2 is a conserved centriole duplication factor. The Journal of cell biology. 188:313-323.

Studier, F.W., and B.A. Moffatt. 1986. Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. Journal of molecular biology. 189:113-130.

Sugino, K., and Y. Naitoh. 1982. Stimulated cross-bridge patterns corresponding to ciliary beating in Paramecium Nature. 295:609-611.

Sullivan, K.F. 1988. Structure and utilization of tubulin isotypes. Annual review of cell biology. 4:687-716.

Summers, K.E., and I.R. Gibbons. 1971. Adenosine triphosphate-induced sliding of tubules in trypsin-treated flagella of sea-urchin sperm. Proceedings of the National Academy of Sciences of the United States of America. 68:3092-3096.

Suryavanshi, S., B. Edde, L.A. Fox, S. Guerrero, R. Hard, T. Hennessey, A. Kabi, D. Malison, D. Pennock, W.S. Sale, D. Wloga, and J. Gaertig. 2010. Tubulin glutamylation regulates ciliary motility by altering inner dynein arm activity. Current biology : CB. 20:435-440.

Takahashi, K., C. Shingyoji, and S. Kamimura. 1982. Microtubule sliding in reactivated flagella. Symposia of the Society for Experimental Biology. 35:159-177.

Tang, C.J., R.H. Fu, K.S. Wu, W.B. Hsu, and T.K. Tang. 2009. CPAP is a cell-cycle regulated protein that controls centriole length. Nature cell biology. 11:825-831.

Tang, W.J., C.W. Bell, W.S. Sale, and I.R. Gibbons. 1982. Structure of the dynein-1 outer arm in sea urchin sperm flagella. I. Analysis by separation of subunits. The Journal of biological chemistry. 257:508-515.

137

Tilney, L.G., J. Bryan, D.J. Bush, K. Fujiwara, M.S. Mooseker, D.B. Murphy, and D.H. Snyder. 1973. Microtubules: evidence for 13 protofilaments. The Journal of cell biology. 59:267-275.

Tokuyasu, K.T. 1975. Dynamics of spermiogenesis in Drosophila melanogaster. VI. Significance of "onion" nebenkern formation. Journal of ultrastructure research. 53:93-112.

Toyoshima, F., S. Matsumura, H. Morimoto, M. Mitsushima, and E. Nishida. 2007. PtdIns(3,4,5)P3 regulates spindle orientation in adherent cells. Developmental cell. 13:796-811.

Toyoshima, Y.Y. 1987a. Chymotryptic digestion of Tetrahymena 22S dynein. I. Decomposition of three-headed 22S dynein to one- and two-headed particles. The Journal of cell biology. 105:887-895.

Toyoshima, Y.Y. 1987b. Chymotryptic digestion of Tetrahymena ciliary dynein. II. Pathway of the degradation of 22S dynein heavy chains. The Journal of cell biology. 105:897- 901.

Tsang, W.Y., C. Bossard, H. Khanna, J. Peranen, A. Swaroop, V. Malhotra, and B.D. Dynlacht. 2008. CP110 suppresses primary cilia formation through its interaction with CEP290, a protein deficient in human ciliary disease. Developmental cell. 15:187- 197.

Uzawa, M., J. Grams, B. Madden, D. Toft, and J.L. Salisbury. 1995. Identification of a complex between centrin and heat shock proteins in CSF-arrested Xenopus oocytes and dissociation of the complex following oocyte activation. Developmental biology. 171:51-59. van Breugel, M., M. Hirono, A. Andreeva, H.A. Yanagisawa, S. Yamaguchi, Y. Nakazawa, N. Morgner, M. Petrovich, I.O. Ebong, C.V. Robinson, C.M. Johnson, D. Veprintsev, and B. Zuber. 2011. Structures of SAS-6 suggest its organization in centrioles. Science. 331:1196-1199.

Veleri, S., S.H. Manjunath, R.N. Fariss, H. May-Simera, M. Brooks, T.A. Foskett, C. Gao, T.A. Longo, P. Liu, K. Nagashima, R.A. Rachel, T. Li, L. Dong, and A. Swaroop. 2014. Ciliopathy-associated gene Cc2d2a promotes assembly of subdistal appendages on the mother centriole during cilia biogenesis. Nature communications. 5:4207.

Venoux, M., X. Tait, R.S. Hames, K.R. Straatman, H.R. Woodland, and A.M. Fry. 2013. Poc1A and Poc1B act together in human cells to ensure centriole integrity. Journal of cell science. 126:163-175.

Vernon, G.G., and D.M. Woolley. 2002. Microtubule displacements at the tips of living flagella. Cell motility and the cytoskeleton. 52:151-160.

Vernon, G.G., and D.M. Woolley. 2004. Basal sliding and the mechanics of oscillation in a mammalian sperm flagellum. Biophysical journal. 87:3934-3944.

Vladar, E.K., and T. Stearns. 2007. Molecular characterization of centriole assembly in ciliated epithelial cells. The Journal of cell biology. 178:31-42.

138

Vonderfecht, T., M.W. Cookson, T.H. Giddings, Jr., C. Clarissa, and M. Winey. 2012. The two human centrin homologues have similar but distinct functions at Tetrahymena basal bodies. Molecular biology of the cell. 23:4766-4777.

Vonderfecht, T., A.J. Stemm-Wolf, M. Hendershott, T.H. Giddings, Jr., J.B. Meehl, and M. Winey. 2011. The two domains of centrin have distinct basal body functions in Tetrahymena. Molecular biology of the cell. 22:2221-2234.

Vorobjev, I.A., and Y.S. Chentsov. 1980. The ultrastructure of centriole in mammalian tissue culture cells. Cell biology international reports. 4:1037-1044.

Vulprecht, J., A. David, A. Tibelius, A. Castiel, G. Konotop, F. Liu, F. Bestvater, M.S. Raab, H. Zentgraf, S. Izraeli, and A. Kramer. 2012. STIL is required for centriole duplication in human cells. Journal of cell science. 125:1353-1362.

Wada, S., M. Okuno, and H. Mohri. 1991. Inner arm dynein ATPase fraction of sea urchin sperm flagella causes active sliding of axonemal outer doublet microtubule. Biochemical and biophysical research communications. 175:173-178.

Wang, Q., A. Suzuki, S. Mariconda, S. Porwollik, and R.M. Harshey. 2005. Sensing wetness: a new role for the bacterial flagellum. The EMBO journal. 24:2034-2042.

Wanner, A., M. Salathe, and T.G. O'Riordan. 1996. Mucociliary clearance in the airways. American journal of respiratory and critical care medicine. 154:1868-1902.

Washburn, M.P., D. Wolters, and J.R. Yates, 3rd. 2001. Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nature biotechnology. 19:242-247.

Weber, K., N. Geisler, U. Plessmann, A. Bremerich, K.F. Lechtreck, and M. Melkonian. 1993. SF-assemblin, the structural protein of the 2-nm filaments from striated microtubule associated fibers of algal flagellar roots, forms a segmented coiled coil. The Journal of cell biology. 121:837-845.

Werner, M.E., P. Hwang, F. Huisman, P. Taborek, C.C. Yu, and B.J. Mitchell. 2011. Actin and microtubules drive differential aspects of planar cell polarity in multiciliated cells. The Journal of cell biology. 195:19-26.

Williams, C.L., C. Li, K. Kida, P.N. Inglis, S. Mohan, L. Semenec, N.J. Bialas, R.M. Stupay, N. Chen, O.E. Blacque, B.K. Yoder, and M.R. Leroux. 2011. MKS and NPHP modules cooperate to establish basal body/transition zone membrane associations and ciliary gate function during ciliogenesis. The Journal of cell biology. 192:1023- 1041.

Williams, N.E., and J. Frankel. 1973. Regulation of microtubules in Tetrahymena. I. Electron microscopy of oral replacement. The Journal of cell biology. 56:441-457.

Williams, N.E., J.E. Honts, and J. Kaczanowska. 1990. The formation of basal body domains in the membrane skeleton of Tetrahymena. Development. 109:935-942.

Williams, N.E., and O.H. Scherbaum. 1959. Morphogenetic events in normal and synchronously dividing Tetrahymena. Journal of embryology and experimental morphology. 7:241-256.

139

Winey, M., A.J. Stemm-Wolf, T.H. Giddings, Jr., and C.G. Pearson. 2012. Cytological analysis of Tetrahymena thermophila. Methods in cell biology. 109:357-378.

Wloga, D., D. Dave, J. Meagley, K. Rogowski, M. Jerka-Dziadosz, and J. Gaertig. 2010. Hyperglutamylation of tubulin can either stabilize or destabilize microtubules in the same cell. Eukaryotic cell. 9:184-193.

Wloga, D., and J. Gaertig. 2010. Post-translational modifications of microtubules. Journal of cell science. 123:3447-3455.

Wloga, D., K. Rogowski, N. Sharma, J. Van Dijk, C. Janke, B. Edde, M.H. Bre, N. Levilliers, V. Redeker, J. Duan, M.A. Gorovsky, M. Jerka-Dziadosz, and J. Gaertig. 2008. Glutamylation on alpha-tubulin is not essential but affects the assembly and functions of a subset of microtubules in Tetrahymena thermophila. Eukaryotic cell. 7:1362-1372.

Wloga, D., D.M. Webster, K. Rogowski, M.H. Bre, N. Levilliers, M. Jerka-Dziadosz, C. Janke, S.T. Dougan, and J. Gaertig. 2009. TTLL3 Is a tubulin glycine ligase that regulates the assembly of cilia. Developmental cell. 16:867-876.

Wolf, N., D. Hirsh, and J.R. McIntosh. 1978. Spermatogenesis in males of the free-living nematode, Caenorhabditis elegans. Journal of ultrastructure research. 63:155-169.

Wolff, A., B. de Nechaud, D. Chillet, H. Mazarguil, E. Desbruyeres, S. Audebert, B. Edde, F. Gros, and P. Denoulet. 1992. Distribution of glutamylated alpha and beta-tubulin in mouse tissues using a specific monoclonal antibody, GT335. European journal of cell biology. 59:425-432.

Woolley, D.M. 2010. Flagellar oscillation: a commentary on proposed mechanisms. Biological reviews of the Cambridge Philosophical Society. 85:453-470.

Woolley, D.M., and G.G. Vernon. 2001. A study of helical and planar waves on sea urchin sperm flagella, with a theory of how they are generated. The Journal of experimental biology. 204:1333-1345.

Xia, L., B. Hai, Y. Gao, D. Burnette, R. Thazhath, J. Duan, M.H. Bre, N. Levilliers, M.A. Gorovsky, and J. Gaertig. 2000. Polyglycylation of tubulin is essential and affects cell motility and division in Tetrahymena thermophila. The Journal of cell biology. 149:1097-1106.

Yan, X., R. Habedanck, and E.A. Nigg. 2006. A complex of two centrosomal proteins, CAP350 and FOP, cooperates with EB1 in microtubule anchoring. Molecular biology of the cell. 17:634-644.

Yang, P., D.R. Diener, C. Yang, T. Kohno, G.J. Pazour, J.M. Dienes, N.S. Agrin, S.M. King, W.S. Sale, R. Kamiya, J.L. Rosenbaum, and G.B. Witman. 2006. Radial spoke proteins of Chlamydomonas flagella. Journal of cell science. 119:1165-1174.

Yano-Toyoshima, Y. 1985. Two heavy chains of 21S dynein from sea urchin sperm flagella. Journal of biochemistry. 98:767-779.

Yano, Y., and T. Miki-Noumura. 1980. Sliding velocity between outer doublet microtubules of sea-urchin sperm axonemes. Journal of cell science. 44:169-186.

140

Yubuki, N., and B.S. Leander. 2013. Evolution of microtubule organizing centers across the tree of eukaryotes. The Plant journal : for cell and molecular biology. 75:230-244.

Zhang, S., and B.J. Mitchell. 2015. Basal bodies in Xenopus. Cilia. 5:2.

Zhao, H., L. Zhu, Y. Zhu, J. Cao, S. Li, Q. Huang, T. Xu, X. Huang, X. Yan, and X. Zhu. 2013. The Cep63 paralogue Deup1 enables massive de novo centriole biogenesis for vertebrate multiciliogenesis. Nature cell biology. 15:1434-1444.

Zou, C., J. Li, Y. Bai, W.T. Gunning, D.E. Wazer, V. Band, and Q. Gao. 2005. Centrobin: a novel daughter centriole-associated protein that is required for centriole duplication. The Journal of cell biology. 171:437-445.

141