The central pair apparatus of the ciliary axoneme is a multi-protein complex important for

proper ciliary beating in motile cilia

Daniel Chen Dai (260563818)

Department of Anatomy and Cell Biology Faculty of Medicine McGill University, Montreal, Canada April, 2020

A thesis submitted to McGill University in partial fulfillment of the requirements of the degree of Master of Science

© Daniel Chen Dai, 2020

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TABLE OF CONTENT

ABSTRACT ...... 4 RÉSUMÉ ...... 6 ACKNOWLEDGMENT ...... 8 ABBREVIATIONS ...... 11 INTRODUCTION ...... 12 RATIONALE, HYPOTHESIS AND AIMS ...... 13 CHAPTER 1 – LITERATURE REVIEW ...... 15 1.1 THE AXONEME: THE CILIARY STRUCTURE ...... 15 1.2 CILIOPATHIES: PRIMARY CILIARY DYSKINESIA ...... 16 1.3 CLINICAL IMPACT AND PATIENT SYMPTOMS ...... 18 1.4 METHODS FOR DIAGNOSING PCD ...... 19 1.4.1 Ultrastructural based diagnosing ...... 19 1.4.2 Ciliary motility-based diagnosing ...... 20 1.4.3 Nasal nitric oxide testing ...... 21 1.4.5 Genetic screening ...... 21 1.5 THE CENTRAL PAIR ...... 22 1.5.1 Structure and function...... 22 1.5.2 Proteins of the CP ...... 24 1.6 CURRENTS METHODS OF STUDYING THE CP PROTEINS...... 28 1.7 FIGURES ...... 30 CHAPTER 2 – IDENTIFICATION AND MAPPING OF CENTRAL PAIR PROTEINS BY PROTEOMIC ANALYSIS ...... 31 2.1 PREFACE ...... 32 2.2 ABSTRACT ...... 33 2.3 INTRODUCTION ...... 34 2.4 MATERIALS AND METHODS ...... 37 Strains and culture condition ...... 37 Chlamydomonas flagella isolation and purification of microtubule fraction ...... 37 Cross-sectional EM ...... 38 Cryo-EM observation ...... 38 Cryo-ET ...... 39 MS analysis...... 39 Data analysis ...... 40 2.5 RESULTS AND DISCUSSION ...... 41 2.5.1 Purification of the axoneme fraction retains the CP proteins with a minimal amount of unrelated proteins ...... 41 2.5.2 Re-evaluation of CP proteins by comparative proteomic analysis ...... 42 2.5.3 Localizing CP protein candidates into sub-structures of the CP complex ...... 46 2.5.4 C1a/e proteins ...... 47 2.5.5 C1a-e-c complex proteins ...... 48 2.5.6 C1d proteins ...... 48 2.5.7 C1c proteins ...... 49 2.5.8 C1b/f proteins ...... 50 2.5.9 C2b proteins ...... 50 2.5.10 Proteins localized at C2a, c, d, e and bridge ...... 51 2.5.11 Other CP protein candidates ...... 52 2.5.12 Model of CP protein localization and insights into functions in the flagella ...... 53 2.6 CONCLUSION ...... 54 2.7 TABLES ...... 56 2.8 FIGURES ...... 60

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2.9 SUPPLEMENTARY MATERIAL ...... 70 2.10 REFERENCES ...... 82 CHAPTER 3 - GENERAL DISCUSSION, SUMMARY AND CONCLUSION ...... 86 3.1 GENERAL DISCUSSION AND SUMMARY ...... 86 3.2 CONCLUSION: ...... 88 3.3 FUTURE PLAN: ...... 89 REFERENCES ...... 90

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ABSTRACT

Cilia or flagella of eukaryotes are small micro-hair-like structures indispensable to single- cell motility and play an important role in mammalian biological processes. Cilia or flagella are composed of nine doublet microtubules surrounding a pair of singlet microtubules called the central pair (CP). Together, this arrangement forms the canonical and highly conserved “9+2” axonemal structure. The CP, which is a unique structure exclusive to motile cilia, is a pair of structurally dimorphic singlet microtubules decorated with numerous associated protein complexes termed C1 and C2. The corresponding protein complexes which bind around their respective microtubule are therefore named C1a to C1f and C2a to C2e. Mutations of CP- associated proteins have been a cause of many different symptoms in cilia related diseases termed ciliopathies. Therefore, there is a necessity to further our understanding of the architecture of the

CP. At this moment there is a distinct lack of knowledge at the most basic level of understanding of the CP. The size of the central pair and its respective sub-complexes in relation to the 22 proteins known to localize there strongly suggest that many more proteins are involved. To this day the proteins that make up the CP remain largely unknown. Our limited understanding of the proteins that comprise the CP is due to the limitations of traditional methods for the identification of CP proteins. These methods relied exclusively on the availability of Chlamydomonas mutants of CP proteins created through random insertional mutagenesis. The central aim of this thesis is to demonstrate a more efficient methodology for identifying new proteins of the CP to provide a full and comprehensive understanding of the proteins at play. To do this, we apply a combination of mass spectrometry (MS) and mutational based analysis in the model organisms Chlamydomonas reinhardtii. The proteomic composition of the entire CP was identified by comparing MS of wild- type (WT) cells and pf15 cells, which are missing the entire CP structure. To map CP proteins into

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specific sub-structures, Chlamydomonas reinhardtii strains of different CP structural mutants were subjected to an identical MS analysis. Mutants missing parts of the CP structure: pf16 (C1- less strain), pf6 (missing C1a complex) and cpc1 (lacking C1b complex), were used to generate

MS profiles for each newly identified CP protein and, afterwards, for mapping into sub-structures based on known proteins. Flagella were purified from WT and mutant cells; pf15, pf16, pf6, and cpc1, treated with 0.6M NaCl twice to remove dynein motor complexes, and analyzed by MS.

MS was performed by in-gel digestion 3 times for each strain and statistical analysis was performed. Comparison between wild type and pf15 MS results revealed over 40 new CP protein candidates in addition to previously well-characterized CP proteins. By comparing these results to subsequent mutant MS results, those proteins were further mapped to the sub-structures surrounding the CP.

Taken together, the CP is a complex structure composed of over 60 CP related proteins that radially arrange themselves around a specific singlet to a specific CP sub-complex. We believe that our approach is faster and less restrictive and has provided further insights into the composition and localization of new proteins of the CP through which future studies can build on.

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RÉSUMÉ

Les cils ou flagelles des eucaryotes sont de petites structures micro-poilues indispensables

à la motilité d'une seule cellule et jouent un rôle important dans les processus biologiques des mammifères. Elles sont composés de neuf microtubules doublets entourant une paire de microtubules simples appelée paire centrale (PC). Ensemble, cet arrangement forme une structure axonémique 9+2 canonique et hautement conservée. La PC, qui est une structure unique exclusive aux cils mobiles, est une paire de microtubules singulets structurellement dimorphiques décorés de nombreux complexes protéiques associés appelés C1 et C2. Les complexes protéiques correspondants qui se lient autour de leur microtubule respective sont donc nommés C1a à C1f et

C2a à C2e. Les mutations des protéines associées à la PC sont à l'origine de nombreux symptômes différents chez les maladies liées aux cils appelées ciliopathies. Il est donc nécessaire d'approfondir notre compréhension de l'architecture de la PC. À l'heure actuelle, il existe un manque flagrant de connaissances au niveau le plus élémentaire de la compréhension de la PC. La taille de la PC et de ses sous-complexes respectifs par rapport aux 22 protéines connues comme y étants localisées suggère fortement que beaucoup plus de protéines sont impliquées. À ce jour, les protéines qui composent la PC restent largement inconnues. Notre compréhension limitée des protéines qui composent la PC est due aux limites des méthodes traditionnelles d'identification des protéines de la PC. En effet ces méthodes reposent exclusivement sur la disponibilité de Chlamydomonas qui sont des proteins de la PC créés par mutagenèse d’insertion aléatoire. L'objectif principal de cette thèse est de présenter une méthodologie plus efficace pour l'identification de nouvelles protéines de la PC afin de fournir une compréhension complète et globale des protéines en jeu. Pour ce faire, nous avons appliqué une combinaison de spectrométrie de masse (MS) et d'analyse basée sur les mutations dans les organismes modèles Chlamydomonas reinhardtii. La composition protéomique

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de la PC entière a été identifiée en comparant la MS des cellules de type sauvage et des cellules pf15, chez qui la structure pc est manquante. Pour cartographier les protéines de la PC en sous- structures spécifiques, des souches de Chlamydomonas reinhardtii de différents mutants structurels de la PC ont été soumises à une analyse MS identique. Les mutants chez qui il manque des portions de la structure de la PC : pf16 (souche sans C1), pf6 (complexe C1a manquant) et cpc1 (complexe C1b manquant), ont été utilisés pour générer le profil MS de chaque protéine de

PC nouvellement identifiée et, par conséquent, pour la cartographie en sous-structures basées sur des protéines connues. Les flagelles ont été purifiées à partir de cellules de type sauvage et mutantes ; pf15, pf16, pf6 et cpc1, traitées avec du NaCl 0,6M deux fois pour éliminer les complexes moteurs de la dynéine, et analysées par MS. Celle-ci ayant ètè realisèe en digestion par gel 3 fois par souche , suivi d’une analyse statistique. La comparaison entre les résultats de la MS de type sauvage et de la MS pf15 a révélé plus de 40 nouvelles protéines PC candidates qui viennent s’ajouter aux protéines PC précédemment bien caractérisées. En comparant ces résultats

à ceux de la MS mutante, ces protéines ont été mises en correspondance avec les sous-structures entourant la PC.

Dans l'ensemble, la PC est une structure complexe composée de plus de 60 protéines liées

à la PC qui s'organisent radialement autour d'un singulet spécifique pour former un sous-complexe spécifique à la PC. Nous pensons que notre approche est plus rapide et moins restrictive et qu'elle a permis de mieux comprendre la composition et la localisation des nouvelles protéines de la PC.

Nous espérons ainsi que ces methodes pourrons être utilisées comme base pour de futures études.

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ACKNOWLEDGMENT

I would first like to recognize my supervisor, Dr. Khanh-Huy Bui whose temperament, humor and unwavering support has been more akin to a close friend than that of a supervisor alone.

I consider myself both honored and lucky at the same time for being a part of the first generation of trainees to graduate from the lab. Without his knowledge and expertise, I would not have been able to come as far as I did and graduate with the pedigree I have now. I am forever indebted to him for helping me become a better person, scientist, and member of society. It will have been an honor to graduate as a student but more importantly a friend.

In addition to my supervisor, I would like to recognize Dr. Muneyoshi Ichikawa. As my first mentor in the lab, he taught me proper science for the first time and what it meant to think like a scientist. All my skills and areas of expertise all derive from his initial teachings. More importantly, I hope to never forget the long conversation we had together about Dr. Bui and the many running jokes and ideas we had. If Dr. Bui guided me through my masters, it was Dr.

Muneyoshi Ichikawa that was the catalyst that helped me decide to do it. My only regret is that you left McGill to become a professor of your own before you could see me graduate.

I would like to thank the amazing team at FEMR, Jeannie Mui, Kelly Sears and Kaustuv

Basu and the members of the MUHC proteomics laboratory, Amy Wong and Lorne Taylor whom without I would have no research or results. Their efforts and expertise were an invaluable asset to me and critical for my projects.

To the members of my committee Dr. Bechstedt, Dr. Ortega and Dr. William Tsang, I would like to thank them for their support and counselling in and outside the lab.

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Finally, I would like to thank all members of the Bui lab and peers past and present whom without them my life these last two years would not have been as colourful.

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CONTRIBUTIONS OF AUTHORS

This dissertation includes one unpublished manuscript of which the master’s candidate is the first author. Contributions of all authors are provided as below:

Identification and mapping of central pair proteins by proteomic analysis.

Daniel Dai, Muneyoshi Ichikawa, Katya Peri, Reid Rebinsky and Khanh-Huy Bui.

The candidates designed and performed all experiments and data analysis, and wrote the manuscript

Muneyoshi Ichikawa helped design, conceive, and supervised the project and wrote the manuscript.

Katya Peri assisted in preforming throughout experiments and proofread the manuscript.

Reid Rebinsky assisted in preforming the experiments and proofread the manuscript.

Khanh-Huy Bui helped design, conceive, and supervise the project.

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ABBREVIATIONS

CP Central Pair

CPC1 Central Pair-Associated Complex 1

DMT Doublet Microtubules

DNAI1 Dynein Axonemal Intermediate Chain 1

FAP Flagellar Associated Protein

IDA Inner Dynein Arms

KLP1 Kinesin-Like Protein 1

MS Mass Spectrometry

Nit1 Nitrate Reductase 1

NO Nitric Oxide

ODA Outer Dynein Arms

PCD Primary Ciliary Dyskinesia

PF Paralyzed Flagella

RS Radial Spoke

TEM Transmission Electron Microscopy

Wild-Type WT

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INTRODUCTION

The evolutionarily conserved cilia and flagella are terms often used synonymously to describe the same hair-like appendage1. It protrudes from the surface of almost all eukaryotic cells. Although there are subtle differences in their length and localization, there are no consistent differences in their structure and function and the term cilia and flagella will be used interchangeably2. Cilia resemble small hairs oriented perpendicularly to the surface of the cell membrane from which they extend. They may vary in length, amount and degree of motility3. The simplest and commonest form of categorization that can be applied to cilia is dependent on its motility or lack thereof. The two forms are referred to most commonly as primary cilia (immotile) and motile cilia4. In its most primordial form, motile cilia can be found on single cellular organisms and are used as a form of propulsion for movements such as in Chlamydomonas reinhardtii or Tetrahymena thermophila5,6.

In higher-level organisms such as humans, motile cilia decorate the surface of our respiratory tract, playing a pivotal role in respiratory maintenance in a process known as mucociliary clearance7.

Well-timed and coordinated beating in both single-cell motility and multicellular systems is a prerequisite to directional movement and proper functioning biological systems8. Primary cilia although similar in composition and form play a much more different role. Primary cilia are integral in many different forms of cell signaling as many different receptors, ion channels, transport proteins, and downstream effectors localize to this cilium9. Forms of sensory stimulation to which primary cilia has been shown to respond to include mechanical stress (bending), chemo- sensation (morphogens) and in some unique cases to light, temperature, osmolality, and gravity10-

13. One such system for primary cilium cell signaling is in the olfactory system. Olfactory sensory neurons that act as the receptor for the olfactory system are bipolar neurons whose dendrites end at the olfactory knob where their primary cilia are located14. Olfactory cilia are responsible for our

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sense of smell15. Within the context of primary cilium cell signaling, odorants contact the olfactory epithelium as a form of ligand initiating olfactory signaling through G-protein-coupled receptors on the surface of olfactory sensory neurons15,16. In stark contrast to motile cilia which can manipulate the surrounding environment to generate gliding or rhythmic movement, primary cilia are subject to their environment to convert external stimuli into a cell signal-dependent response17.

Cilia decorates the surface of all eukaryotic cells in one form or another and therefore have an extended area of influence on many different systems from single-cell motility to more complex biological systems. Due to its ubiquitous distribution throughout the human body, complications in proper cilia function can be directly linked to several ciliopathies (cilia related diseases) such as

Bardet Biedl syndrome and Primary Ciliary Dyskinesia (PCD)18. On top of that many ciliopathies exhibit a heterogeneous nature. Multiple different defects or mutations often lead to the same disease with multiple causative genes potentially responsible. Therefore, there is an urgency to better understand cilia and flagella to identify as many proteins as possible in order to address system specific variants.

Rationale, hypothesis and aims

Rationale: Due to the heterogeneous nature of PCD and other ciliopathies many patients are diagnosed late in life with many patients perhaps not being diagnosed at all. The current forms of diagnosis are directly limited by our knowledge of the axoneme and its structure8 (Fig. 1.7.1). One such gap in our knowledge is the proteins that comprise the central pair (CP), a pair of singlet microtubules found at the center of motile cilia. Research has shown that mutations at the CP often lead to motility defects in model organisms19,20. Clinical evidence has shown that mutations of the

CP proteins in patients later confirmed in Chlamydomonas reinhardtii do not produce easily identifiable morphological defects yet retain the potency to cause ciliopathies equivalent to motor

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protein defects21. The CP, however, remains poorly understood at both a functional and compositional level. The current methods in studying CP proteins are thorough and complete, however, due to the randomness of mutant generations it remains slow and inefficient to meet the current demand. Our group has applied mass spectrometry of purified axoneme from wild-type

(WT) Chlamydomonas reinhardtii to address this hurdle. Through comparative MS with purified axoneme from specialized axonemal mutant made available by the Chlamydomonas library we compare WT MS data of structural mutants of the CP22. Through this analysis we are not only able to identify most CP proteins very quickly, but we are also able to begin roughly localizing them around the central pair. By identifying as many CP proteins as possible we can expedite functional and structural studies at the CP which can directly impact the success of diagnosis at the clinical level as well as open new avenues to possible gene therapy targets.

Hypothesis – The CP is a large protein complex that is home to more proteins than currently known. Mass spectrometry is a fast and effective means of identifying these new proteins of the central pair proteome with high accuracy.

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CHAPTER 1 – LITERATURE REVIEW

1.1 The Axoneme: the ciliary structure

Cilia and flagella are conserved organelles whose function and structure are shared among all cilia bearing organisms1. Both primary and motile cilia are comprised of nine radially arranged specialized microtubules called doublet microtubules (DMT)23. The DMT consist of one 13 protofilament (composed of alternating α- and β-tubulin monomers) singlet microtubule called the

A-tubule with an incomplete 10 protofilament microtubule-binding onto it called the B-tubule.

This specific arrangement of nine DMTs has been termed the axoneme and plays host to hundreds of proteins related to structure and function (Fig. 1.7.1)23. One of the major underlying differences between primary and motile cilia is whether the cilia have an inherent ability to generate movement. This difference can be observed at the functional and structural levels. The axoneme of primary cilia when observed in cross-section consists of nine radially arranged DMTs or more traditionally referred to as the canonical ”9+0” arrangement such as olfactory cilium4. However, when compared to motile cilia, several important proteins and structures become exclusive. First and foremost is the presence of motor protein complex dynein at the outer (outer dynein arms,

ODA) and inner (inner dynein arms, IDA) periphery of each DMT23. These motor proteins help to facilitate microtubule sliding between adjacent DMTs and therefore movement in an ATP dependent fashion24. These dynein arms are large protein complexes, each comprised of several heavy, intermediate and light chains25. More notably is the presence of an additional microtubule- based structure in motile cilia. At the center of the nine DMTs are two structurally dimorphic 13 protofilament singlet microtubules bound together by protein complexes26. Like the DMTs, this complex, termed the central pair, is also home to many more additional axonemal protein and protein complexes26. The axoneme of motile cilia is therefore considered to be organized in a

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“9+2” arrangement such as in the case of respiratory cilia27 (Fig. 1.7.1). It is worth noting, however, that although these are the commonly held understanding, several special cases do exist. Nodal cilia are an important factor in determining left/right symmetry in developing embryos but despite being motile it lacks the CP and is, therefore, a form of motile “9+0” cilia28. The reverse is true in that kinocilium located in the sensory epithelium of vertebrate inner ears are a form of non-motile cilia with a CP and are therefore a form of non-motile “9+2” cilia14. Radial spoke (RS) proteins can also be found on the surface of DMTs. They protrude towards the center of the axoneme23.

This multiprotein complex is essential in regulating dynein arm activity and relaying signals from the CP to the motor arms29. More recently a whole new family of proteins was discovered to bind within the lumen of each DMT called microtubule inner proteins although their functions have yet to be deduced30. These proteins all together help to facilitate a waveform like motion in order to generate movement24. Motile cilia are able to beat at a frequency ranging from approximately 8-

20 Hz under basal condition but can be accelerated by contact with an irritant or intracellular calcium fluxes31,32. Taken all together, the axoneme is an extremely densely populated and complex structure with specific complexes and proteins unique to primary and motile cilia. Due to the conservation of the axoneme across the phylogenetic spectrum, proteins such as dynein, RS and the CP structure remain conserved. We are therefore able to use the biflagellate single cellular green alga Chlamydomonas reinhardtii as a model for motile cilia.

1.2 Ciliopathies: Primary Ciliary Dyskinesia

There is an increasing amount of diseases stemming from dysfunction in the axoneme of motile cilia. These types of diseases are collectively known as ciliopathies. In the specific case of motile cilia, ciliopathies often describe conditions where motile cilia do not beat properly or at all33. Several of these diseases include PCD or Kartagener syndrome8. In diseases such as PCD,

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motile cilia of the respiratory tract have impaired mucociliary clearance which results in several clinical manifestations, including increased susceptibility to chronic bronchitis leading to bronchiectasis, chronic rhinosinusitis, chronic otitis media, situs inversus totalis, and male infertility8. PCD has been characterized as a rare genetically heterogeneous disorder inherited as an autosomal recessive disorder with an incidence rate ranging from one in 12,000 to 17,0007,33.

This means that between 1700 and 2500 Canadians could find themselves afflicted with this disease. Due to it being genetically heterogeneous, pin-pointing a causative gene or genes has been a difficult and a daunting challenge through conventional family-based genome-wide linkage studies18. To compensate for the limitations of conventional techniques, a combination of different methods has been successfully employed in order to identify PCD related genes. These methods include functional candidate gene testing, homozygosity mapping, comparative genomics, and proteomics34-37. Together several genes have been identified and linked to PCD, many of which belong to the dynein motor protein complex8. Proteins such as dynein axonemal intermediate chain

1 (DNAI1) were among the first proteins identified through candidate gene testing38. These findings were validated after a study of 179 PCD patients where 16 of these patients held biallelic mutations for DNA1 in addition to 12 other novel mutations39. What is perhaps even more surprising was the discovery of mutations in RS proteins of the axoneme which are not directly involved in motor-driven cilia movement. Both RS head protein 9 and 4A were identified through homozygosity mapping in families diagnosed with PCD40. In the case of both dynein and RS proteins, mutations in the biflagellate organism Chlamydomonas reinhardtii corresponding to their respective human orthologs produced visible defects. At the ODA, IDA and RS complex these mutations resulted in gross ultrastructural abnormalities8. The resulting mutation in

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Chlamydomonas reinhardtii leads to a loss of swimming, consistent with cilia dysfunction in PCD patients29,41,42.

1.3 Clinical impact and patient symptoms

In the human system, motile ciliated cells can be found primarily lining the walls of the respiratory tract from the paranasal sinuses to the terminal airways43. Together the cilia beat in synchrony to move periciliary fluid and mucus from the lower respiratory tract otherwise known as the mucociliary escalator44. The mucociliary escalator are one of the primary defense mechanisms of the respiratory system and any functional aberration is strongly linked to respiratory and pulmonary complications7. Clinical manifestations of PCD can be identified as early as the neonatal period. Over 70-80% of patients may show signs of respiratory distress during this early stage8. This strongly suggests that motile cilia at an early stage are critical for fetal lung fluid clearance. In some cases, newborns may even suffer from acute respiratory failure, hypoxemia, and pneumonia, and require immediate mechanical ventilation8. A common occurrence in almost half the cases of PCD is situs inversus totalis which is a complete left-right mirroring of the internal organs18. Without properly functioning nodal cilia, the left-right laterality appears to be random. Patients with organ misplacement in the form of situs inversus totalis or in milder forms such as a singular organ laterality defects are at a higher risk for congenital cardiovascular defects45. In almost all cases of diagnosed PCD, patients have reported experiencing rhinosinusitis (inflammation of the nasal passage) as well as otitis media (middle ear disease)8. Both symptoms have been described in varying degrees in almost all PCD cases and appear to be the primary features of the disease. Otitis media may lead to hearing loss later in life46.

A prevalent symptom of PCD is a persistent cough. This is to compensate for the lack of a mucociliary escalator meant to clear the lower respiratory tract47. If the lower respiratory tract is

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unable to clear mucous, bacterial cultures may persist leading to more frequent episodes of pneumonia, bronchitis and bacterial-induced infections7. Bronchiectasis (inflammation of the walls of the lungs respiratory tract) follows multiple bacterial infections. Bronchiectasis has a direct effect on respiration in the form of hypoxemia and persistent breathing exacerbation episodes. In the later stages of a patients’ life, it may even become necessary for a lung transplant in order to reduce the likelihood of death.48. Adverse effects on the pulmonary system overall have a major impact to the quality of life of patients as they get older49. Male specific symptoms in addition to those mentioned before may include impaired sperm motility leading to impotence, but this is not true in all cases50. In females, the effect of PCD on the motile cilia of the fallopian tubes are still unclear but appears to have no detrimental effect on their reproductive organs8.

1.4 Methods for diagnosing PCD

One of the greatest hurdles in PCD treatment is an immediate and accurate diagnosis of the disease7. Awareness of PCD and ciliopathies are still growing in the public and clinical eye. Often early symptoms of PCD are misdiagnosed or attributed to other diseases with similar symptoms.

Otitis media or middle ear disease is a very common symptom among PCD patients; however, it is not uncommon that a patient is instead referred to an otolaryngologist. At this point, a successful diagnosis is fully dependent on the otolaryngologist suspicion for PCD8. As a result, most PCD cases are often diagnosed later in life or left undiagnosed altogether. PCD diagnosing is an evolving field as currently there is no standardized diagnostic procedure. More and more methods for identifying the disease are currently being tested.

1.4.1 Ultrastructural based diagnosing

The most classical means of testing for PCD to this day is a thorough ultrastructural analysis of patient motile cilia axoneme through traditional transmission electron microscopy

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(TEM)51. In this method, the overall structure of the axoneme in cross-section is examined for the presence of any obvious defects. As mentioned before common defects include missing densities corresponding to any of the dynein motor protein, RS proteins or the CP in its entirety. There are many different types of axonemal malformities but many of them are likely to be unrelated to PCD and can be false positives attributed to structural defects related to infection and inflammation52.

The major distinction between these secondary forms of structural defects is a heterogeneous distribution of these defects, whereas defects associated with PCD seem to be present in almost all afflicted motile cilia. In perfectly healthy individuals with no signs of PCD, around 5% of cilia in the system can be considered defective in both function and structure53. TEM is thus an effective means for diagnosing PCD in obvious cases. The limitations of these methods, however, rely completely on known disease-causing mutations that produce an ultrastructural defect in the axoneme. Unfortunately, in most cases patients with clear signs of PCD will have a completely intact and regular looking axonemes54. In addition, this type of testing is not easily accessible.

Hospitals or research centers may not be equipped with specially tailored microscopes and technicians in order to conduct these tests furthering the need to find alternative methods for diagnosing.

1.4.2 Ciliary motility-based diagnosing

An obvious area for examination is whether sperm or respiratory cilia are motile. Typically, what is a good indication of PCD is the presence of completely paralyzed flagella thus resembling primary cilia. Another degree of immobilization includes a dyskinetic ciliary activity. This is easily scene in the respiratory pathways where synchronous beating is required. Asynchronous beating relative to neighboring cilia is usually accompanied by an attenuated ciliary beat frequency of around three Hz or lower at room temperature. There are recorded cases of the opposite effect

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occurring in PCD patient and therefore further research in ciliary kinetics is needed to improve this means of diagnosis8.

1.4.3 Nasal nitric oxide testing

Nasal nitric oxide (NO) level has recently developed as a form of non-invasive means for identifying PCD. Recent studies have shown that nasal NO levels are significantly reduced when compared to healthy patients55. The reason and mechanism for NO levels and its relationship to motile cilia have yet to be deduced.

1.4.5 Genetic screening

Genetic screening as a means for identifying PCD may show the most promise and accuracy moving forwards. Whole exome sequencing has been used in order to identify mutations in known PCD genes. The success of a genetic screen, however, is directly related to the number of known PCD genes. Although this number is ever-increasing, not enough is known in order to compensate for the heterogeneous nature of PCD. Known PCD genes that are often looked for are contained in the dynein motor complexes (DNAI1, DNAH5, etc.)38,56. These mutations are often easily identifiable in an ultrastructural analysis as well. Unfortunately, only a small fraction of patients will harbor mutations in these genes. The effectiveness of diagnosing through genetic screening is greatly affected by our knowledge of the proteins and their function at the axoneme.

Currently, knowledge on the proteins of the axoneme is in its infancy with structures such as the

CP remaining relatively unknown.

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1.5 The central pair

1.5.1 Structure and function

The CP is a unique microtubule-based structure present at the center of nine DMTs forming the canonical “9+2” arrangement in motile cilia. It consists of two structurally dimorphic singlet microtubules termed C1 and C2, respectively. Each 13-protofilament singlet microtubule is decorated with a variety of unique protein complexes. These protein complexes are named based on the singlet they bind onto, thus, C1a to C1f and C2a to C2e bind to the C1 and C2 microtubule respectively26. The two singlets are bound together by a large protein complexes called the bridge and diagonal link57 (Fig. 1.7.2a). All together the function and role the CP plays appear to be diverse and complex specific.

The role of the CP has yet to be fully deduced however many studies have reported multiple different effects, thus painting the CP as a multi-faceted structure. One thought is that the CP is a regulator of ciliary movement. Oddly enough the CP is not exclusively linked to any other structure surrounding it and appears to have a free 360° rotation within the axoneme itself58. Movement of cilia is dynein dependent, however, the force generated by dynein is only able to influence the

DMT adjacent to it59. Due to the radial symmetrical distribution, activation of dynein on one end will be canceled by the same force generated on the opposite side. This dilemma caused by the radial symmetry of the dyneins requires differential regulation of the motor mechanism in order to produce net movement58,60. The CP is thought to be this regulator for several reasons. First, the orientation of the CP seems to be correlated to the direction of the beat. The CP is often orientated at right angles to the ciliary power stroke in a variety of metazoan cilia61. Secondly, many mutants in Chlamydomonas at the CP with obvious missing protein structures or tubules are paralyzed even though their dynein function remains intact19,21,57,62-64. Finally, the relationship between the CP

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and the RS may be critical in signaling and regulating movement, although this relationship has not been fully defined. Mutations in RS proteins have shown to also result in paralyzed flagella8,29.

Rotation in the CP was originally observed in Paramecium through TEM images of the axoneme in cross-section. Beating cilia were “instantaneously fixed” from Paramecium with there cilia fixed at different phases of their beating. 20 to 30 serial sections of a singular cilium along the axoneme were observed. From these serial images, the rotation of the CP was tracked using its unique asymmetric protein complexes as morphological markers. In relation to DMTs, it was revealed that the protein complexes were rotating along with the CP58. Similar experiments were done on other organisms including Chlamydomonas. The force that drives CP rotation is still unclear however several hypotheses have been put forward. One possible scenario is an enzymatic activity of the motor protein kinesin found to be located at the CP. Through kinesin, the CP could be actively rotated. Another hypothesis suggests that the CP is being passively rotated by the bending of the axoneme. In solution, the CP adopts a helical conformation60,65. When a bend is propagated throughout the axoneme the helical shape may orient itself to conform to the bend in order to achieve the lowest possible energy conformation to accommodate the generated force60.

Rotation of the CP may help to facilitate differential regulation of the dynein motors through specific interactions with select RS complex in its proximity. The CP is ideally situated to regulate the sliding of the outer DMTs. Through a rotational dependent regulation, the CP may explain the diversity in different waveforms and patterns found in different organisms despite sharing the same

“9+2” structure60. This is consistent with nodal cilia a CP-less variant of motile cilia exhibiting only a singular rotational waveform rather than any complex two- and three-dimensional waveforms found in the “9+2” motile cilia66.

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1.5.2 Proteins of the CP

The CP is likely comprised of many more proteins then there are currently reported. Of the

22 known proteins that localize at the CP many of them have specific roles that may serve to fine- tune the ciliary waveform.

1.5.2.1 Central pair-associated complex 1

Central pair-associated complex 1 (CPC1) was among one of the first structurally significant proteins identified and localized to the CP. The CPC1 deficient strain was created randomly through insertional mutagenesis of a nitrate reductase 1 (Nit1) cassette into Nit1 deficient

Chlamydomonas67. Subsequent screening for swimming phenotypes was used to identify potentially relevant mutations. Structurally the CPC1 strain of Chlamydomonas axoneme is malformed at the C1b protein complex. In fact, in the absence of CPC1 protein, the C1b complex does not appear to assemble at the CP suggesting that it may be a prerequisite for proper assembly or localization of the C1b complex (Fig. 1.7.2a). CPC1 deficient strains of Chlamydomonas beat asymmetrically and have a reduced beat frequency of about two thirds when compared to WT while still retaining a regular waveform57. The cDNA corresponding to CPC1 revealed two predicted functional domains, a single EF-hand motif at the C-terminus and a centrally located adenylate kinase domain. It is thought that the long narrow arrangement of the axoneme may restrict the supply of ATP into the flagellar compartments by diffusion64. Thus, there must be some mechanisms through which ATP concentration is maintained in order to activate dynein sufficiently. This reasoning is consistent with reduced beat frequencies in CPC1 deficient

Chlamydomonas. It was later shown that saturating amounts of ATP could partially rescue CPC1 associated defects in vitro64. Therefore, CPC1 may be a local ATP regulator specifically at the location of the C1b complex of the CP in order to maintain proper dynein motor activity. Because,

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cilia frequency is only reduced and not fully paralyzed, this suggests there must be other forms of

ATP regulation occurring either at the CP or in its surroundings. The presence of an EF-hand motif suggests another possible level of regulation. There may be a form of calcium-dependent regulation on CPC1 activity however, this hypothesis has not yet been fully explored.

1.5.2.2 Paralyzed flagella 6 and calmodulin

Paralyzed flagella (PF) 6 was obtained from random insertional mutagenesis of Nit1 in

Nit1 Chlamydomonas and screened for motility defects like CPC1. Motility defects in pf6 contained strains that jiggled or twitched with no net progress68. Ultrastructural analysis of the axoneme revealed that the C1a complex of the CP was missing entirely (Fig. 1.7.2b). Sequence analysis of the PF6 gene indicates that it encodes for a large polypeptide with many alanine-, proline-rich and basic domains. No obvious functions could be deduced from the sequence alone.

Both the phenotype and the structural defect could be rescued by phage clone cotransformation of a full-length copy of the PF6 gene. This observation suggests that the protein encoded by PF6 is both important for the assembly of the C1a complex and that the C1a complex is regulating ciliary motility differently then the C1b complex based on the phenotype observed. Using antibodies against PF6, a combination of immunoprecipitation and MS was used to identify the amino acid identities of five polypeptides that could form a complex with PF6, presumably, the C1a complex.

Among these polypeptides able to form a complex with PF6 was calmodulin, an intermediate calcium-binding messenger protein63. The presence of calmodulin has several implications relating to dynein regulation. It was shown previously that although dynein activity is largely ATP based, dynein activity could be increased in a calcium-dependent manner69. Dynein in mutant strains of

Chlamydomonas lacking the C1 microtubule entirely in addition to the C1a was immune to the effects of calcium induction70. PF6 may be important for the assembly of the C1a complex and the

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recruitment of calmodulin63. The presence of a calcium-sensing protein at the C1a complex suggest a calcium-dependent form of motility tuning present at the CP at the location of the C1a complex.

1.5.2.3 Paralyzed flagella 16

PF16 mutants were generated randomly using Nit1 insertional mutagenesis. The resulting structural phenotype in PF16 strains is a structurally unstable C1 microtubule. Many cross- sectional images of the PF16 axoneme contain partial to completely missing C1 microtubule and all its surface binding protein complexes57 (Fig. 1.7.2b). Sequence analysis of PF16 reveals that the protein encoded contained several repeating alpha-helices otherwise characteristic of several armadillo repeats whose role and function are diverse in many armadillo repeat containing proteins62. Antibodies raised against a fusion protein expressed from the cloned cDNA of PF16 were used to localize PF16 along the length of WT flagella. A combination of immunogold labeling and TEM demonstrated that the C1 microtubule was specifically being decorated with gold beads62. Motility in C1 compromised PF16 mutants was completely paralyzed. The mechanisms in which PF16 is involved in C1 stabilization and where the protein specifically resides at the C1 microtubule remains unclear.

1.5.2.4 Kinesin-like protein 1

Kinesin-like protein 1 (KLP1) encodes for a phosphoprotein of the kinesin superfamily. It was found to be present at the C2 microtubule at the C2c and C2b regions through immunogold labeling. Like most conventional kinesins it binds to microtubules in vitro in the presence of adenosine 5′-[β,γ-imido]triphosphate instead of ATP71. At the position of the C2c complex KLP1 is in an optimal place to interact with the radial spoke proteins of the DMTs71. RNAi knockdown of KLP1 in Chlamydomonas exhibit reduced swimming. In some cases, it was reduced to half the

WT beat frequency71.

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1.5.2.5 Hydin, a potential source of clinical ambiguities

As mentioned before there isn’t a single diagnostic test able to accurately identify PCD but rather, there are multiple tests acting complementary to one another to achieve this goal. In a structural analysis of PCD patients, physicians rely strongly on the presence of universally identifiable gross structural defects. Common defects being completely missing projections or structures such as missing dynein, RS or the entire CP in rare cases. Any sign of these defects often may lead to an accurate diagnosis. However, in most cases, patients appear to have completely normal axonemes meaning the dynein, RS and CP are all still present54. In these scenarios what might be possibly occurring is a mutation at the axonemal level that cannot be resolved at the resolution of standard TEM. The CP, in this case may, be a potential source for these mutations.

Not much is quite known about the CP however mutations at the CP represent a grey area in structural analysis. Hydin is a CP protein whose effects were previously identified in mice with congenital hydrocephalus72. Through homozygosity mapping of three PCD afflicted siblings, a mutation in the gene HYDIN was identified. After whole-exome wide sequencing, it was revealed that they all held homozygous nonsense mutations of the gene HYDIN. However, in these patients, most of their axonemal cross-sections appeared normal21. It was later shown in a mutational assay in Chlamydomonas that the protein encoded by HYDIN was able to produce a visible morphological defect important for the assembly of the C2b projection. Cilia activity in HYDIN deficient Chlamydomonas showed very weak to no activity consistent with PCD patient cilia19,73.

It is very unlikely that HYDIN is an isolated case and that many CP proteins are critical for proper ciliary function but do not produce easily identifiable mutation observable under standard TEM methods in humans. Most mutational studies in other organisms at the central pair report defects in ciliary motility, thus the CP represents a genetic hotspot for mutations to occur which are

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directly linked to cilia motility. Hydin is a now well-characterized CP protein and PCD causing gene. It is thought to be located at the C2b complex on the CP19. Hydin serves as evidence that not all mutations at the CP manifest itself at the structural level but have similar impacts at the clinical level in humans. The current problem facing physicians now is three-fold. First, due to the heterogeneity of ciliopathies such as PCD, multiple proteins in the axoneme may be a causative gene. Secondly, many mutations or defects in ciliary proteins produce little to no morphological defects as seen in many patients8. Finally, the CP, a unique structure found mostly in motile cilia remains to be fully understood in both function and composition. To address challenges at the clinical level a basic and fundamental understanding of the CP and the proteins that comprise it is an essential first step that this thesis attempts to take.

1.6 Currents methods of studying the CP proteins

Despite an ever-increasing need to explore the central pair proteome the methods employed to do so have remained slow and inefficient. The ciliary axoneme of Chlamydomonas is composed of over more than 700 proteins20. The CP is a large and complex structure at the axoneme yet only

22 proteins have been identified at this location. This inconsistency is one of the primary motivations in our study of the CP proteome. Currently, the study of CP proteins is primarily conducted in the model organism Chlamydomonas reinhardtii. Groups that study CP proteins rely mainly on random mutagenesis through either UV or random DNA insertions into the

Chlamydomonas genome in order to generate mutations22,67,74. Following this, a mutation must be able to produce a mutant phenotype and more importantly a morphological defect at the central pair before further analysis continues. This workflow has been a gold standard for identifying structurally significant CP proteins to date including potentially causative ciliopathic genes19.

Through this method, both protein and function are deduced through a single workflow. The

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bottleneck, however, is the random nature of mutagenesis. Many research groups continue to rely on random mutations generated in-lab or purchased from the Chlamydomonas library where hundreds of mutations are generated randomly and stored22. The issue remains however that each mutation generated must be characterized. This involves multiple rounds of genetic backcrossing to remove any unwanted background mutations as a result of random mutagenesis. An extensive number of motility-based assays, measurements and rescue experiments is required to assess any phenotype differences. Most importantly is a thorough ultrastructural analysis of the axoneme in cross-section with TEM to identify any defects. The time it takes, however, to begin this from start to end in order to identify a single CP protein is grossly inefficient. Many mutants are studied in parallel in order to compensate for the randomness of CP mutant generation. Thus, there is a clear necessity for a full and comprehensive analysis of the CP proteome in order to expedite the functional and structural assays that need to follow. I thus propose in this thesis that the use of mass spectrometry as a comprehensive technique for identifying new CP proteins both fast and accurately.

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1.7 Figures

Figure 1.7.1: Motile cilia axoneme in cross-section. A schematic diagram of the axoneme of the canonical “9+2” structure of motile cilia and major axonemal proteins.

Figure 1.7.2: CP schematic and select CP mutants. (A) A schematic diagram of the CP in cross- section from WT axoneme adapted from Carbajal-González et al., 201326 . (B) schematic diagram of the structural mutants pf16, cpc1 and pf6 and their observed defects57,62-64.

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CHAPTER 2 – Identification and mapping of central pair proteins by

proteomic analysis

Daniel Dai1, Muneyoshi Ichikawa2, Katya Peri1, Reid Rebinsky1 Khanh Huy Bui1

1 Department of Anatomy and Cell Biology, McGill University, Montréal, Québec H3A 0C7.

Canada

2 Department of Systems Biology, Graduate School of Biological Sciences, Nara Institute of

Science and Technology, 8916-5, Takayama-cho, Ikoma, Nara 630-0192, Japan

Corresponding authors:

Muneyoshi Ichikawa, Department of Systems Biology, Graduate School of Biological Sciences,

Nara Institute of Science and Technology, 8916-5, Takayama-cho, Ikoma, Nara 630-0192, Japan. e-mail: [email protected]

Khanh Huy Bui, Department of Anatomy and Cell Biology, McGill University, Montréal, Québec

H3A 0C7. Canada. e-mail: [email protected]

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2.1 Preface

In this chapter we identify the proteins of the CP using MS on WT and compare them to the MS of four unique CP ultrastructural mutants in order to roughly localize them around the CP.

The goal is to use MS as a principal technique in order to rapidly identify new proteins that belong to the central pair. Through a comparative proteomic analysis with wild type and CP structural mutants we have identified 40 additional proteins belonging to the central pair and their potential locations around the CP. Through this methodology we have accelerated the rate at which new CP proteins were being identified. This allows future studies to circumvent the need for random mutagenesis and phenotype screening to identify potentially new CP proteins. By identifying most if not all the proteins of the CP we have expedited the speed at which CP proteins can be studied at, thus, allowing for more impactful structural and functional studies which may help to improve the clinical battle against ciliopathies.

Very recently another group studying the CP published a study during the time when we were preparing our manuscript for this study. Zhao et al (2019)., also relied on MS as the primary methods for identifying proteins of the CP. Both groups were able to identify similar proteins thus validating the technique. However, in our study we were able to map and localize most of our newly identified CP proteins with MS. This allowed for a more comprehensive and detailed model of the arrangement, relationship and potential interactions of the proteins of the CP.

The study in this chapter has been accepted and is awaiting publication in the journal

Biophysics and Physicobiology published by The Biophysical Society of Japan ©.

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2.2 Abstract

Cilia or flagella of eukaryotes are small micro-hair like structures that are indispensable to single-cell motility and play an important role in mammalian biological processes. Cilia or flagella are composed of nine doublet microtubules surrounding a pair of singlet microtubules called the central pair (CP). Together, this arrangement forms a canonical and highly conserved 9+2 axonemal structure. The CP, which is a unique structure exclusive to motile cilia, is a pair of structurally dimorphic singlet microtubules decorated with numerous associated proteins.

Mutations of CP-associated proteins cause several different physical symptoms termed as ciliopathies. Thus, it is crucial to understand the architecture of the CP. However, the protein composition of the CP was poorly understood. This was because the traditional method of identification of CP proteins was mostly limited by available Chlamydomonas mutants of CP proteins. Recently, more CP protein candidates were presented based on mass spectrometry results, but most of these proteins were not validated. In this study, we re-evaluated the CP proteins by conducting a similar comprehensive CP proteome analysis comparing the mass spectrometry results of the axoneme sample prepared from Chlamydomonas strains with and without CP complex. We identified a similar set of CP protein candidates and additional new 11 CP protein candidates. Furthermore, by using Chlamydomonas strains lacking specific CP sub-structures, we present a more complete model of localization for these CP proteins. This work has established a new foundation for understanding the function of the CP complex in future studies.

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2.3 Introduction

Cilia and flagella are common terms used to describe the same hair-like structure of eukaryotic cells and will therefore be used interchangeably in this paper. It is known that defective cilia are implicated in a variety of different human diseases, from developmental disorders to metabolic syndromes 1. However not all cilia are alike. Primary cilia are non-motile and are commonly reported as sensory receptors 2. Motile cilia, on the other hand, show a beating motion at high frequencies driven by motor protein dynein 3. This rudimentary motion is the driving force for a plethora of multi-level systems from single cell movement to mammalian organ function and maintenance 1. Motile cilia are also present in our respiratory system and beat together in order to clear mucus build up and infectious agents 4.

Cilia-related diseases, otherwise known as ciliopathies, such as primary ciliary dyskinesia

(PCD), are derived from the impairment of motile cilia 5,6. Patients who suffer from PCD often experience a wide spectrum of symptoms ranging from male infertility to an increased susceptibility to respiratory infections 7. Failure to recognize or diagnose PCD early on can often be lethal later in life 4. A common practice used to diagnose PCD is a cross-section analysis of patient nasal epithelium cilia using transmission electron microscopy (TEM). However due to the diversity of PCD mutations, many different defective proteins can lead to similar malformations

7. Additionally, not all mutations produce visible differences at standard TEM resolution while still inducing PCD-like symptoms. The largest obstacle to our understanding of cilia-related defects is our limited comprehension of the proteins that comprise cilia.

Cilia are highly complex structures composed of different compartments. Motile cilia consist of nine doublet microtubules (DMTs) surrounding a pair of singlet microtubules called the central pair (CP) 8. This specific arrangement defines the “9+2” structure of the axoneme (Fig.

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2.8.1A). There exists axonemal dyneins (outer dynein arm, ODA, and inner dynein arm, IDA) attached to DMTs which are responsible for the beating of cilia. Radial spoke (RS) complexes extend from DMTs toward the CP. Intraflagellar transport (IFT) driven by IFT dyneins and IFT kinesins takes place on the DMTs 9. This arrangement of the axoneme structure is highly conserved in all eukaryotes with motile cilia, suggesting that there exists a similar set of proteins and processes. Thus, we can study the axoneme composition using model organisms like the green algae Chlamydomonas reinhardtii.

The presence of the CP distinguishes motile cilia from its immotile counterpart, primary cilia. The CP has a diverse array of functions including the regulation of local Ca2+ concentration,

ATP/ADP concentration and axonemal dynein activities through mechanical interactions with the

RS 10-13. The CP is a huge protein complex composed of a pair of structurally and functionally dimorphic singlet microtubules named C1 and C2 and many other associated proteins 14. The CP has a variety of sub-structures such as C1a, C1b, C1c and C1d on the C1 singlet, or C2a, C2b and

C2c on the C2 singlet that were characterized by traditional cross-sectional electron microscopy

(EM). C1 and C2 microtubules are connected by two structures called the “bridge” and the

“diagonal link”. With recent higher resolution cryo-electron tomography (cryo-ET) structures, more details of these sub-structures have been characterized allowing the C1a to be sub-classified as C1a/e, the C1b as C1b/f, the C2a as C2a/e and the C2c as C2c/d (Fig. 2.8.1B) 15. Through this manuscript, we follow the newer nomenclature of the CP sub-structures as in Fig. 2.8.1B. These sub-structures bind with 16- or 32-nm repeating units along the axonemal axis 15. Despite their unique existence in motile cilia and their importance to motility, the proteins that comprise the CP have been poorly understood. Traditionally, 22 proteins (apart from α- and β-tubulins) have been characterized as components of the CP complex (Table 2.7.1). For example, kinesin-like protein 1

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(KLP1) is a phosphoprotein that localizes at the C2 microtubule around C2c/d region 16. Mutations in several known CP proteins such as Hydin located at the C2b and FAP221 (PCDP1) located at the C1d have been previously shown to cause ciliopathic symptoms 10,17. However, it was generally believed that there should be more unidentified proteins inside the CP complex. Due to random insertions of transgenes into the Chlamydomonas reinhardtii genome, previous characterization of

CP proteins largely relied on phenotype-based screening of CP protein mutants 18,19. This approach, however, remains inefficient and biased towards proteins which produce visible phenotypes. Recently, Zhao et al. used a more comprehensive approach taking advantage of relative quantitative mass spectrometry (MS) comparing Chlamydomonas strains with and without intact CP 20. By doing so, 44 proteins were proposed to be new CP protein candidates in addition to the traditionally known CP proteins. Though some of these proteins, FAP47, FAP76, FAP99,

FAP246 and DPY30, were confirmed to locate at the CP by immunofluorescence study, the other candidates remain to be tested. Furthermore, the localizations of these new CP protein candidates in CP sub-structures were not clear.

Here, we re-examined the CP protein components by a similar proteomic approach and obtained mostly the same result but with 11 additional CP protein candidates. Furthermore, we localized these newer CP protein candidates to certain CP sub-structures by using several different

Chlamydomonas mutant strains of CP sub-structures. Our results have established a new foundation for understanding the CP architecture.

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2.4 Materials and Methods

Strains and culture condition

Chlamydomonas reinhardtii strains used in this study are as follows: CC-124 (Wild Type, WT),

CC-1033 (pf15, CP-less mutant) 21, CC-5148 (cpc1, C1b/f-less mutant) 14, CC-1034 (pf16, mutant with an unstable C1 structure) 14,22,23 and CC-1029 (pf6, C1a/e-less mutant) 15,24. The cells were purchased from the Chlamydomonas resource center. Cells were grown first on a Tris- acetatephosphate (TAP) 25 solid plate containing 1.5% agar and then cultured in TAP liquid media under shaking or stirring conditions under a 12 hr light and dark cycle. For flagella purification, each Chlamydomonas strain c was cultured in 1 L liquid TAP media with stirring until the OD600 reached 0.5-0.6.

Chlamydomonas flagella isolation and purification of microtubule fraction

The cells were harvested by low-speed centrifugation (700g for 7 min at 4℃), and flagella were removed from the cell bodies by pH shock 26. Cell bodies were removed by low speed centrifugation (1,800g for 5 min at 4℃) in HMDS (10mM HEPES, pH7.4, 5mM MgSO4, 1mM

DTT, 4% sucrose containing 10 μg/ml aprotinin and 5 μg/ml leupeptin) and flagella were collected by higher-speed centrifugation (4,696g for 40 min at 4℃). Isolated flagella were resuspended in

HMDEKP buffer (30 mM HEPES, pH 7.4, 5 mM MgSO4, 1 mM DTT, 0.5 mM EGTA, 25 mM potassium acetate, 0.5% polyethylene glycol, (MW 20,000) containing 10 μM paclitaxel, 1 mM

Phenylmethylsulfonyl fluoride (PMSF), 10 μg/ml aprotinin and 5 μg/ml leupeptin). Paclitaxel,

PMSF, leupeptin and aprotinin were added to the buffer throughout the purification after this step.

Flagella were demembraned by incubating with HMDEKP buffer containing 1.5% NP40 for 30 min on ice. For cryo-electron microscopy (cryo-EM), sonication was performed for better splitting of the axoneme after NP40 treatment. Chlamydomonas axonemes were then spun down by a

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tabletop centrifuge (7,800g for 10 min at 4℃). To split the bundle of axonemes, the samples were incubated with ADP (final concentration of 1 mM) for 10 min at room temperature to activate dyneins and then incubated with 0.1 mM ATP for 10 min at room temperature to induce doublet sliding. The samples were then spun down (16,000g for 10 min at 4℃). Protease was not added for splitting. After this, Chlamydomonas microtubule fraction was incubated twice with HMDEKP buffer containing 0.6 M NaCl for 30 min on ice, spun down (16,000g for 10 min at 4℃) and resuspended in HMDEKP buffer. Purification process was performed three times for each strain for biological triplicates.

Cross-sectional EM

For cross-sectional EM, the samples (200-300 μg/ml) were fixed overnight at 4°C in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) and washed three times with washing buffer (0.1 M sodium cacodylate buffer). Samples were post fixed with 1% aqueous OsO4 and

1.5% aqueous potassium ferrocyanide for 2 h and washed three times with washing buffer.

Specimens were dehydrated in a graded ethanol-dH2O series 30, 50, 70, 80, 90 to 100% ethanol.

The samples were infiltrated with a graded Epon-ethanol series (1:1, 3:1), embedded in 100% Epon and then polymerized in an oven at 60°C for 48 hr. Ultrathin sections (90 to 100-nm thickness) were prepared from the polymerized blocks with a Diatome diamond knife using a Leica

Microsystems EM UC7 ultramicrotome, transferred onto 200-mesh copper grids and stained with

4% uranyl acetate for 6 min and Reynold’s lead for 5 min. The TEM grids were imaged by a FEI

Tecnai G2 Spirit 120 kV TEM equipped with a Gatan Ultrascan 4000 CCD Camera Model 895.

Cryo-EM observation

3.5 μl of sonicated microtubule fraction sample (~4 mg/ml) purified from WT Chlamydomonas was applied to glow-discharged holey carbon grids (Quantifoil R2/2), blotted and vitrified in

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liquid ethane using the Vitrobot Mark IV (Thermo Fisher Scientific). Micrographs were obtained at 59kx nominal magnification on the direct electron detector Falcon II with the Titan

Krios (Thermo Fisher Scientific) using a total dose of ~28 electrons/Å2 and 7 frames (calibrated pixel size of 1.375 Å/pixel). The defocus range was set to between −1.2 and −3.8 μm.

Cryo-ET

A demembranated WT Chlamydomonas flagella sample (~500 μg/ml, 3.5 μl) was applied to glow-discharged holey carbon grids (Quantifoil R2/2), blotted and vitrified in liquid ethane using the Vitrobot Mark IV (Thermo Fisher Scientific). Vitrified grids were then loaded onto

Titan Krios (Thermo Fisher Scientific) equipped with Falcon II direct electron detector (Thermo

Fisher Scientific) operating at 300 kV. Tilt series were obtained using tomography software

(Thermo Fisher Scientific) in the range of -40 to +60 degrees with a 4-degree increment at nominal magnification of 29kx so that the total dose was limited to ~100 electrons/Å2. The defocus was set to −5 μm.

MS analysis

4x Laemmli buffer (#1610747, Bio-Rad) was added to the microtubule fraction samples in

HMDEKP buffer so that it will be 1x, and 25-30 µg protein was loaded onto the SDS-PAGE gel.

To detect all proteins in the sample, electrophoresis was terminated before the proteins entered the separation gel. A single band containing all proteins in the sample was then cut out from the gel and subjected to in-gel digestion 27. Resultant ~2 μg peptides, which is the maximum protein amount that can be loaded, were chromatographically separated on a Dionex Ultimate 3000

UHPLC. First, peptides were loaded onto a Thermo Acclaim Pepmap (Thermo, 75 µm ID × 2 cm with 3 µm C18 beads) precolumn and then onto an Acclaim Pepmap Easyspray (Thermo, 75 µm

× 25 cm with 2 µm C18 beads) analytical column and separated with a flow rate of 200 nl/min

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with a gradient of 2-35% solvent (acetonitrile containing 0.1% formic acid) over 2 hours.

Peptides of charge 2+ or higher were recorded using a Thermo Orbitrap Fusion mass spectrometer operating at 120,000 resolution (FWHM in MS1, 15,000 for MS/MS). The data was searched against Chlamydomonas reinhardtii protein dataset from UniProt (https://www.uniprot.org/).

Data analysis

MS data was analyzed using Scaffold_4.8.4 (Proteome Software Inc.). Total spectrum counts of each protein was divided by total spectrum of the whole sample for normalization to obtain quantitative values. Proteins with mean values of exclusive unique peptide counts of 2 or more in the WT MS results were used for further analysis. To identify CP protein candidates, Student’s t- test was applied to pf15 and WT MS results using biological triplicates. Proteins exhibiting a minimum four-fold change and a statistical significance threshold (p < 0.05) in pf15 or proteins which were completely missing in pf15 were categorized as CP candidates. For statistical analysis using several mutant strain MS results, one-way analysis of variance (ANOVA) followed by

Dunnett’s multiple comparisons test was performed using GraphPad Prism 8.

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2.5 Results and Discussion

2.5.1 Purification of the axoneme fraction retains the CP proteins with a minimal amount of unrelated proteins

Previous approaches targeting each CP protein one by one presented clear evidence for 22 proteins10,14,17,21,23,24,28-30 (Table 2.7.1). A recent study comparing MS results from the

Chlamydomonas axoneme with and without a whole CP complex presented more candidates of

CP proteins but most of them were not validated 20. Here, we aimed to re-evaluate the CP protein composition by similar MS analysis. Due to the sensitive nature of MS, peptide detection tends to have an unfavorable preference for large and abundant proteins. Previous proteomic analysis of whole Chlamydomonas flagella showed the presence of an abundant amount of proteins from membrane and matrix fractions and large proteins such as dynein heavy chains 20,31. Therefore, we tried to obtain samples which contain CP proteins with less contaminant proteins. To achieve this, the microtubule fraction was purified from WT Chlamydomonas flagella by sequential purification following the deflagellation by pH shock. First, proteins from the membrane and matrix fraction were removed by NP-40 treatment. Flagella and demembranated axoneme samples were verified by cross-sectional EM and cryo-ET. The axoneme containing two singlet microtubules belonging to the CP with protruding sub-structures were observed (Fig. 2.8.1C and Fig. S2.9.1A).

Demembranated axonemes were then treated with ADP/ATP and with 0.6 M NaCl twice to extract axonemal dyneins (Fig. 2.8.2A). From SDS-PAGE, significant amounts of proteins were removed in the final extract leaving the tubulin band which is a main component of the CP and DMT (Fig.

2.8.2B). Though we tried to remove the RS complexes by dialysis against a low salt buffer 32 or a

KI treatment 33, it was not possible to remove Chlamydomonas RS complexes while maintaining the integrity of the microtubule structure. Thus, the RS complexes remained in our sample. It was

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previously reported that a high salt treatment destabilizes the CP structure, especially the C2 microtubule 14. Cross-sectional EM result of our NaCl treated sample were consistent with this

(Fig. S2.9.1B). Nevertheless, twice salt treated sample was also imaged using cryo-EM and the remaining singlet microtubules from the CP were confirmed to retain the repeating sub-structures

(Fig. 2.8.2C and Fig. S2.9.2).

To detect all proteins in the sample, the purified microtubule fraction was subjected to

SDS-PAGE, and proteins were cut out as a single band and analyzed by MS (see details for

Materials and Methods). In our MS results, almost all (21 out of 22) known CP proteins such as

Hydin, CPC1, PF6, FAP69, PF16, KLP1 and FAP221 (PCDP1) were detected by our MS (Table

2.7.1). The only known CP protein which we failed to detect was FAP227 (C1a-18) 34. Since the size of FAP227 is small (18 kDa), it is thought to be unfavorable for detection by MS. Detected traditionally known CP proteins were previously shown to localize to different CP sub-structures

(Table 2.7.1). Together with the EM results, we concluded that our purification method retained the CP proteins and was usable for proteomic analysis.

Known microtubule inner proteins (MIPs) like Rib43a and Rib72 35,36 and RS proteins were also detected since these structures are tightly associated with DMTs (Table S2.9.1). Proteins from other axonemal components such as IFT complex proteins, IFT dyneins, IFT kinesins, axonemal dyneins and dynein regulatory complex were still detected due to the high sensitivity of MS though we tried to reduce them as much as possible.

2.5.2 Re-evaluation of CP proteins by comparative proteomic analysis

Next, we tried to identify the CP proteins by MS. Since it is not possible to sub-fractionate

CPs from DMTs as they are both microtubule-based structures, we used a comparative proteomic approach using a specific Chlamydomonas mutant lacking a whole CP complex similar to Zhao et

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al., (2019) 20. Chlamydomonas mutant strain pf15 contains a mutation in a gene encoding the p80 subunit of microtubule severing enzyme Katanin 21. The resulting effects prevent the entire CP complex from assembling and lead to paralyzed flagella while leaving other components like

DMTs intact (Fig. 2.8.3A). pf15 strain was chosen in this study instead of pf18 used in 20 since the genetic background of the p15 strain has been well-characterized. An identical purification process was used for pf15 mutant flagella and WT (Fig. 2.8.3A and B). The MS analysis was performed for the microtubule fraction of pf15, and normalized MS results of pf15 and WT were compared

(see details for Materials and Methods). In the MS result of pf15, there were many proteins significantly reduced or completely missing when compared with the WT (Fig. 2.8.3C, left side).

Along with traditionally known CP proteins listed in Table 2.7.1, there were proteins totally missing or significantly reduced in pf15. These proteins were thought to be candidates CP proteins.

There were also proteins detected in higher amounts in pf15 or only detected in pf15 (Fig.

2.8.3C, right side). The numbers of these proteins were less than the proteins that were only detected in WT. In our MS, some IFT complex proteins (IFT22, IFT56, IFT57, IFT74 and BBS1) were detected in higher amounts in pf15 (Table S2.9.1). Previous studies have shown that the IFT complex proteins accumulate in the CP-less Chlamydomonas mutant axonemes, possibly trapped in the empty space where CP complex is supposed to be 37. Therefore, these increased amounts of

IFT proteins are thought to be due to the accumulation of the IFT particles. Other increased proteins might also be trapped inside the axoneme similarly. There were also several proteins only detected in pf15 result, but the amounts of detected peptides of these proteins were generally low, and thus these proteins are thought to be due to contamination (Table S2.9.1).

To clearly see the differences between WT and pf15, we performed a comparison based on protein categories which include tubulins, RS proteins, IFT complex, IFT dynein, IFT kinesin,

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axonemal dyneins, dynein regulatory complex proteins, MIPs and traditionally known CP proteins

(Fig. 2.8.3D). Since values were normalized, α- and β-tubulins were detected at the same level between WT and pf15 results and served as a control. As clusters, only traditionally known CP proteins were significantly decreased in pf15, thus validating our strategy. Other classes like IFT dynein, IFT kinesin, RS proteins, axonemal dyneins, dynein regulatory complex proteins and MIPs did not show any significant decrease as groups. From these results, we concluded that our method is valid to identify CP proteins.

When we focused on each protein, 18 out of the 20 traditionally known CP proteins detected in WT were completely missing or significantly decreased in pf15 (Table 2.7.1). The exceptions were Calmodulin and FAP174 which did not show significant decrease and fell into the region between two-fold decrease and two-fold increase (between two orange lines in Fig.

2.8.3D). Calmodulin is shared with the RS complex 33, and thus its decrease is thought to be masked by signals from the remaining RS in our purification. FAP174 also did not show significant decrease, and this could mean that FAP174 is also present in other axonemal structures (discussed later). Other CP proteins like enolase and HSP70, which are also known to be shared with other axonemal components, showed significant decrease but fell into the region between two- to four- fold decreases in pf15 (between lower orange and blue lines in Fig. 2.8.3D). This area also contained proteins such as DHC9, p38 and KAP from other categories. Almost all the proteins from other complexes did not show decreases of four-fold or larger except DHC8 (below the lower blue line in Fig. 2.8.3D). Thus, proteins which were decreased significantly (p < 0.05) with a 4- fold decrease or greater in pf15, or completely missing in pf15, were categorized as new CP protein candidates. With this criteria, 37 proteins including FAP7, FAP47, FAP65, FAP70 were identified as CP candidates (Fig. 2.8.3D and Table 2.7.2).

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A recent proteomic study used CP-less mutant pf18 instead of pf15 to identify CP proteins

20. Only demembranated whole axonemes were analyzed by quantitative MS in 20 instead of purified microtubule fractions. Despite the differences in strains and methods used, 26 out of 37 identified proteins in our work were shared with Zhao et al., (2019) 20 making these proteins strong candidates for the CP proteins (asterisks in Table 2.7.2). One of these shared CP protein candidates was FAP125. FAP125 is a kinesin-like protein which was previously proposed to be part of the

CP complex without direct evidence 38. Our study presented further evidence showing that FAP125 is a CP-associated kinesin in addition to KLP-1.

Eleven CP candidate proteins were uniquely identified in our study while 19 where unique to Zhao et al. (2019) 20 (Table 2.7.2 and Table S2.9.2). These differences might be due to contaminated proteins from remaining axonemal components in Zhao et al.’s purification method since only demembranated samples were used in the study. Alternatively, it is possible that some of the CP proteins fell off in our purification method. NAP was shown to be a component of the

C1a/e region by immunoprecipitation by Zhao et al., 20, but we failed to detect NAP in our MS.

This could be due to the weak association of NAP to the C1a/e region. It is also possible that differences are due to different strains used. Nevertheless, it is noteworthy that while using different strains and methods, many common proteins were identified as new CP protein candidates. Our work can be therefore be used complementarily to understand the architecture of the CP complex.

One of the proteins identified only in our result was phosphoglycerate mutase which was detected in WT but completely missing in pf15 (Table 2.7.2). Phosphoglycerate mutase was previously shown to be present in the membrane and matrix fraction and play roles in ATP production together with enolase 39. Interestingly, enolase which is involved in the same ATP

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synthesis pathway as phosphoglycerate mutase is known to be a component of the CP as well as being present in the membrane and matrix fractions. Since these enzymes work together, it is likely that phosphoglycerate mutase is also integrated into the CP complex to facilitate the reaction.

In our criteria, FAP244 was also identified as a CP protein. FAP244 was previously shown to be a component of the tether complex of I1 dynein of the IDA by cryo-ET studies 40,41. In our

NaCl treated conditions, FAP244 was detected in WT with high abundancies but not detected in any of the pf15 triplicate (Table 2.7.2). FAP244 was also decreased in the pf18 strain replicates by

Zhao et al., (XP_001694394.1 in Table S2.9.1 in 20) but it was not proposed as a CP protein. It is likely that the FAP244 portion from axonemal dyneins was masking the decrease of FAP244 from the CP complex in the previous study. Based on these results, FAP244 is thought to be shared with the CP complex and the I1 dynein. FAP43 was previously shown to have a weak sequence similarity with FAP244 and to have a redundant function with FAP244 in I1 dynein 40,41. In our

MS results, FAP43 was not decreased in pf15 triplicate (Table S2.9.2). FAP44, which makes up the tether complex of I1 dynein together with FAP43 and FAP243, also did not show any reduction in pf15 (Table S2.9.2). Therefore, FAP244 in the CP complex is thought to have a different and independent function from I1 dynein.

2.5.3 Localizing CP protein candidates into sub-structures of the CP complex

There are several Chlamydomonas mutant strains corresponding to specific CP sub- structures. The pf16 strain is one of such strains with an unstable C1 microtubule 14,22,23. In the pf16 axoneme, the C1 microtubule can be assembled. However, the whole C1 structure is reported to be lost after demembranation or NaCl treatment due to the missing of PF16 protein which localizes to the C1 microtubule 14. Using this feature, Zhao et al., assigned their CP candidates into the C1 or C2 microtubule by comparing MS results of pf16 with that of WT 20. In this study, we

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further aimed to identify the localizations of these CP protein candidates around the CP complex.

To achieve this, we used different kinds of Chlamydomonas mutants lacking specific CP sub- substructures, pf6 (C1a/e mutant), cpc1 (C1b/f mutant) as well as pf16 (Fig. 2.8.4A). In our purification condition, the majority of flagella purified from pf16 strain had only one of the CP microtubules, presumably the C2 microtubule (Fig. S2.9.1C and E) consistent with previous reports. Following the same sample preparations as WT, purified microtubule fractions from different CP mutants were analyzed by MS (Fig. S2.9.3). The normalized MS results from five different strains were compared, and MS detection profiles for each protein of interest were generated (Fig. 2.8.4B-I and Fig. S2.9.4). Traditionally known CP proteins, which localize at the same area, shared similar MS profiles which allowed us to assign newer CP candidates to certain

CP sub-structures.

2.5.4 C1a/e proteins

CP proteins like PF6, FAP101, FAP114 (C1a-32) and FAP119 (C1a-34), which were shown to be located at the C1a/e sub-structure 24,34, were detected both in WT and cpc1 strains since they retain the C1a/e structure. These proteins were either not detected or detected in low amounts in pf6, pf15 and pf16 because of the lack of the C1a/e region (Fig. 2.8.4B). To verify this statistically, we performed one-way ANOVA test and these known C1a/e proteins were all significantly decreased in pf6, pf15 and pf16 but not in cpc1 (Fig. 2.8.4B). DPY30 which was proposed to be located at the C1a/e region by immunoprecipitation 20 also met this standard, further verifying our strategy. Among new CP candidates, FAP7, FAP348 and CHLREDRAFT_150638 showed the same pattern by one-way ANOVA test and were thought to be present in the C1a/e region as well. FAP348 was previously proposed to be located somewhere in the C1 complex 20.

Based on our MS results, FAP348 is likely to be located at C1a/e region. FAP7 was shared with

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Zhao et al.,’s CP candidates but it was not assigned to any region of the CP complex. Our results assigned FAP7 to the C1a/e region. The new CP protein CHLREDRAFT_150638 only identified in our result is also thought to be a component of C1a/e region. After we submitted this manuscript,

FAP216 was localized around the base of C1a/e structure by cryo-ET 42. Though FAP216 showed similar MS pattern with other C1a/e proteins, it was also decreased in cpc1. This could mean that some part of FAP216 is reaching to the cpc1 region or interacting with proteins that compose it.

2.5.5 C1a-e-c complex proteins

Through cryo-ET and immunoprecipitation using PF16 protein as a bait, proteins like

FAP76, FAP81, FAP92, FAP105, FAP108, FAP216, FAP412 and MOT17 (FAP305) were proposed to form a super-complex inside the CP structure 42. In our MS results, most of these proteins (PF16, FAP76, FAP81, FAP92, FAP105 and FAP108) showed similar MS profiles being highly detected in WT, pf6 and cpc1 strains but not detected or detected only very little amount in pf16 and pf15 strains (Fig. 2.8.4D). In terms of the location, some of these proteins like PF16 and

FAP92 are at the C1a/e region 42, but they are thought to be stably tethered to the C1 microtubule due to their interaction even with the apparent loss of the structure in pf6. In our MS analysis,

FAP279 and FAP289 also showed similar MS profiles and were thought to be components of the

C1a-e-d super-complex (Fig. 2.8.4D). FAP279 is a leucine-rich repeat-containing protein that was not assigned as a CP protein before. A homologue of FAP279 is also present in humans (LRRC72).

FAP289 was not previously assigned to certain CP sub-structure.

2.5.6 C1d proteins

There are several CP proteins known to localize at the C1d area. FAP46, FAP54, FAP74,

FAP221 (PCDP1) are such proteins 10. These proteins were proposed to form a complex located at the C1d region. In our MS profile, these proteins show a similar trend being significantly reduced

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only in pf15 and pf16 strains since they lack this region (Fig. 2.8.4E). The difference with known

C1a-c-e complex proteins was that these C1d proteins were still detected in pf16. FAP99 showed a similar MS profile as these C1d proteins and was therefore assigned to this area (Fig. 2.8.4E).

FAP99 was previously confirmed to be an authentic CP protein and was proposed to be located at the C1 microtubule 20. We were then able to further narrow down its localization.

FAP297 (WDR93) was also proposed to be a component of the C1d complex 28 but older work failed to detect this protein 10. Unlike the other C1d proteins, FAP297 showed a significant reduction in the cpc1 strain lacking the C1b/f region (Fig. 2.8.4A). This could mean that FAP297 is located at the interface between the C1d and the C1f and interacting with proteins from the C1f.

Zhao et al., also failed to detect FAP297 by immunoprecipitation using FAP46 as a bait, though all other known C1d proteins were detected 20. This further supports our idea that FAP297 is located away from other known C1d proteins.

2.5.7 C1c proteins

Some proteins like MOT17, FAP125, FAP209 and FAP219 showed a significant reduction only in pf15 and pf16 strains by one-way ANOVA tests. However, these proteins were slightly but not significantly reduced in pf6 (Fig. 2.8.4C). These MS patterns were somewhat in between that of known C1a/e proteins (significant reduction in pf6 result) and C1d proteins (WT level detection in pf6 result). Hence, these proteins are thought to be at the interface of the C1a/e and the C1d, namely the C1c area. One of these proteins, MOT17 was shown to interact with known C1a/e proteins by immunoprecipitation which is the neighboring region of the C1c 20. FAP125 was previously proposed to be somewhere in the C1 microtubule, but its localization to a specific sub- structure was not achieved. Our results located FAP125 into specific sub-structure of the C1 microtubule. For FAP209 and 219, there was no localization information in the axoneme at all.

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2.5.8 C1b/f proteins

Traditionally known proteins belonging to the C1b/f sub-complex like CPC1, FAP42 (C1b-350) and FAP69 (C1b-135) also share similar patterns distinct with significant decrease in cpc1, pf15 and pf16 but with modest decreases in pf16 (Fig. 2.8.4F). At first, we were puzzled with this result of only a modest decrease in pf16 since pf16 is generally assumed to have an unstable, and therefore missing C1 (after a single salt wash) to which the C1b/f region is also attached. Therefore, we looked into the article characterizing the pf16 mutant and realized that the C1b/f part remains with the C2 microtubule due to the diagonal link connecting these structures even though other sub-structures like the C1a, c, d and e, and C2b were missing 14. This was also mentioned by the authors but has been overlooked in recent articles. Thus, we concluded that the C1b/f proteins are partially present in the pf16 structure in our purification condition (Fig. 2.8.4A). Enolase and

HSP70A showed different MS patterns from other known C1b/f proteins (Fig. 2.8.4F), but these proteins were previously shown to be present both in CP complex and in other compartments of the axoneme. The differences are thought to represent the presence of these proteins in other compartments of the axoneme 39,43. Therefore, the MS profile shared with CPC1, FAP42 and

FAP69 was used as a standard for C1b/f proteins. FAP246 was previously shown to interact with other C1b/f proteins by immunoprecipitation 20. Our assignment is consistent with this. Among our new CP candidates, CHLREDRAFT_170023 and CHLREDRAFT_177061 showed C1b/f-like profile and are thought to be located at this region (Fig. 2.8.4F). CHLREDRAFT_170023 and

CHLREDRAFT_177061 were not proposed to be localized to the CP complex before.

2.5.9 C2b proteins

Hydin is the only protein known to be associated with the C2b region 17,30. Based on previous cross-sectional EM results, this region solubilizes before the C1b/f region in pf16 CP

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structure 14. Consistent with this, Hydin was missing in pf16 MS (Fig. 2.8.4G). Hydin was also decreased in cpc1 compared with WT. From previous studies, the C2b projection is in close proximity to the C1b sub-structure 15. It is possible that the interaction between neighboring sub- structures C2b and C1b are tighter than previously assumed. In our MS profile, FAP47 showed a similar trend with Hydin (Fig. 2.8.4G). In recent MS results, FAP47 also showed an elution pattern similar to Hydin 20. Taken together, FAP47 is thought to be located at C2b region. In Zhao et al.,

(2019), FAP49, 72, 154 and 416 were also identified as CP proteins and proposed to form a complex with FAP47 based on their immunoprecipitation results 20. In our MS result, FAP49, 72 and 154 shared peptide sequences and therefore we were not able to conclude if these proteins are decreased in pf15 (Table S2.9.2). FAP416 was not detected in our MS.

2.5.10 Proteins localized at C2a, c, d, e and bridge

KLP1 is known to localize at the C2c/d area 16. KLP1 was detected in most strains but in a very small amounts in the pf15 strain (Fig. 2.8.4H). This is consistent with the results of previous cross-section EM showing that the C2c/d region is stably bound to the C2 microtubule in pf16 14

(Fig. 2.8.4A). Interestingly, PF20 protein which is known to be localized at the “bridge” that connects the C1 and C2 microtubules 44 also showed an MS profile similar with that of KLP1 being detected slightly more in pf16 strain compared with pf15 result (Fig. 2.8.4H). By immunogold labeling EM in previous studies, gold particles were found to be bound to only one of the CP singlets, presumably the C2 microtubule 44. Our results also suggest that PF20 is associated more tightly to the C2 microtubule. Proteins like FAP65, 70, 75, 123, 147, 171, 194, 225, 239, 244 and

266 showed similar MS profiles with KLP1 and PF20 (Fig. 2.8.4H) and therefore are thought to be present at the C2a, c, d, e and the bridge region. The C2a/e area is also included since this part is known to be stably attached to the C2 microtubule in pf16 strain 14. Previous cryo-ET studies

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have shown that there are protein structures inside the C2 microtubule tubulin lattice similar to the

MIPs in the DMTs 15. Some of these proteins might correspond to these inner proteins of the C2 microtubule. FAP65, 70, 75, 147, 171 and 239 were previously proposed to be somewhere at the

C2 microtubule 20 and we were able to further narrow down the localizations of these proteins. For

FAP123, 194, 225 and 266, there was no previous information about their localization inside the

CP complex. FAP244 is thought to be present in both the tether complex of I1 dynein and this region of the CP as mentioned earlier.

2.5.11 Other CP protein candidates

In our results, some of the proteins showed MS profiles which were not readily assigned to certain classes. Calmodulin, a traditionally known CP protein, did not show a significant decrease in either of the strains used in our study (Fig. 2.8.4I). As mentioned, Calmodulin is known to be shared with the RS which was left in our sample preparation. Thus, the decrease of

Calmodulin is thought to be masked by the signal from the RS.

Similarly, FAP174, which is a traditionally known CP protein, did not show significant decrease in either of our strains by one-way ANOVA test (Fig. 2.8.4I). FAP174 might be shared with other compartments of the axoneme similar to Calmodulin. In the previous study, FAP174 was shown to be reduced in CP-less mutant but the signals were still detected by Wester blots 45.

The remaining FAP174 might correspond to the portion from other axonemal components.

Previously, FAP174 was immunoprecipitated by FAP246 with other traditionally known C1b/f proteins 20. In our MS profile, though it was not significant, FAP174 showed a slight decrease in cpc1 which lacks the C1b/f region. Based on these results, FAP174 is thought to be located at the

C1b/f area.

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There were some remaining CP candidates which we were unable to assign to a certain CP sub-structure (Fig. S2.9.4), including FAP312 and 412 which were shared with 20. FAP312 was proposed to be a C2 protein 20 and FAP412 was recently proposed to be a part of C1a-e-c super- complex 42. However, due to low detection of these proteins in our MS, we could not confidently assign these proteins to a certain CP sub-structure. A known C1 protein, PP1a was detected only in WT (Fig. S2.9.4) and therefore we were not able to assign it to a certain sub-structure of the CP.

Though it was not clear by our MS profile (Fig. S2.9.4), phosphoglycerate mutase might be a part of the C1b/f region since this protein is known to work with enolase which was described to localize to this sub-structure. The other CP candidates like EF-3, FAP173 and FAP199 were not assigned to certain sub-structures. These proteins were, however, detected in only small amounts and could be false positives of MS and require further testing.

2.5.12 Model of CP protein localization and insights into functions in the flagella

From our MS profile, we were able to build a more complete model of the localizations of the CP proteins (Fig. 2.8.5 and Table 2.7.2). Our model of localization also gave some insights into the regulation of flagellar motility. FAP125 is a kinesin-like protein recently identified as a

CP protein and our work localized it to the C1c area by our study. The presence of FAP125 at the

C1c area is interesting since KLP-1, another known CP kinesin, is located at the C2c/d which is at the opposite side of the CP complex. KLP-1 was proposed to work as a conformational switch in the CP 16, and thus symmetrical binding of two CP-kinesins onto separate singlet microtubules might play a role in waveform switching or coordinating a planar waveform. Further functional analysis of FAP125 in future work will reveal this point. Like this example, our model of the localizations of CP proteins can be used to understand how each CP protein is organized and is working as a complex.

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Very recently, CP proteins at the C1a-e-c regions were shown to form a super-complex across the CP sub-structures 42. Our MS results also have shown that the components of this super- complex share very similar MS profiles. Other than this region, FAP147 showed a very similar

MS profile with FAP244. FAP266’s MS profile was especially similar with KLP1. Therefore, these protein pairs might also be forming sub-complexes inside the CP structure (Fig. 2.8.5).

Biochemical experiments and structural analysis of the CP complex in a higher resolution would reveal this point in future studies.

2.6 Conclusion

PCD is a rare but prevalent congenital disease that derives from the impairment of motile cilia. An insufficient understanding of the protein composition of axonemal complexes directly affects the success and efficiency of clinical diagnosis of a wide variety of ciliopathies. By using comprehensive MS analysis of Chlamydomonas strains, we have identified more CP proteins and localized them to specific sub-structures of the CP which allows for more informed interpretation of whole exome sequence data and cross-sectional analysis. Through this method, we circumvent traditional means of protein identification and localization and provide a more comprehensive insight into the entire making of the CP complex. Such proteomic approaches that exploit mutant strains could also be applicable for other uninvestigated areas of the axoneme. Our assignments of these CP proteins into each CP sub-structure will serve as primers for future modeling of CP proteins when a high-resolution structure of the CP complex is obtained by cryo-EM.

Acknowledgements

We thank Dr. Kaustuv Basu and Ms. Jeannie Mui at the Facility for Electron Microscopy Research of McGill University for help in preparation of cross-section EM microscope operation and data collection. We are indebted to Mr. Lorne Taylor and Ms. Amy Ho Yee Wong from MUHC for |54|

help with MS. This work was supported by JSPS KAKENHI Grant Number JP19K23726 to MI and grants from the Natural Sciences and Engineering Research Council of Canada (Discovery

Grant 69462), Canada Institute of Health Research (Project Grant 388933) and the Canada Institute for Advanced Research Arzieli Global Scholars Program and McGill University to KHB.

Conflicts of Interest

The authors declare no conflicts of interest.

Author Contributions

MI and KHB conceived the project and designed the experiments. DD and MI performed culture of the cells, purification of microtubule fractions from flagella for MS analysis with the help of

KP and RR. DD performed cross-sectional EM, and MI performed cryo-EM and cryo-ET observations with the aid of KHB. DD and MI analyzed the results. All authors were involved in the manuscript writing process.

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2.7 Tables

Table 2.7.1 Summary of MS results of traditionally known CP proteins.

pf15 WT Exclusive pf15/WT Exclusive unique unique ratio (%) Names Uniprot peptide count p-values peptide count (Quantitative Localizations References ID (Quantitative (WT vs pf15) (Quantitative values were values after values after used) normalization) normalization)

Hydin A8HQ52 68, 87, 27 0, 0, 0 0.0 0.00059 C2b 17,30

(68, 75, 53) (0, 0, 0)

FAP42 A8J614 63, 59, 36 2, 2, 2 3.6 0.00062 C1b/f 39

(C1b-350) (68, 50, 69) (2, 2, 2)

PF6 Q9ATK5 53, 65, 32 4, 3, 5 6.0 0.00016 C1a/e 34,46

(65, 72, 82) (5, 3, 5)

CPC1 Q6J4H1 55, 57, 30 6, 7, 4 8.8 < 0.00010 C1b/f 34

(76, 66, 74) (7, 8, 4)

FAP54 A8J666 46, 61, 27 0, 0, 1 0.7 < 0.00010 C1d 10,28

(44, 48, 49) (0, 0, 1)

FAP46 A8ICS9 40, 50, 35 1, 0, 2 2.5 0.0052 C1d 10,28

(52, 44, 78) (1, 0, 3)

FAP74 D4P3R7 32, 42, 15 0, 0, 1 1.0 < 0.00010 C1d 10,28

(34, 32, 33) (0, 0, 1)

FAP69 A8IF19 16, 24, 11 1, 2, 2 8.3 0.0023 C1b/f 39

(C1b-135) (17, 23, 26) (1, 2, 2)

PF16 A8J0A5 17, 19, 16 1, 1, 1 1.9 0.010 C1 23

(34, 69, 74) (1, 1, 1) C1a-e-c complex 42

HSP70† A8JEU4 14, 22, 6 5, 5, 2 33 0.028 C1b/f 39,43

(17, 18, 10) (7, 6, 2)

KLP1 A8I9T2 15, 16, 3 1, 2, 1 15 0.025 C2c/d 16,47

(17, 13, 7) (1, 2, 2)

FAP101 A8I345 13, 16, 14 0, 0, 1 1.6 0.0055 C1a/e 34

(17, 20, 30) (0, 0, 1)

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Enolase† A8JH98 17, 12, 9 7, 9, 6 39 0.017 C1b/f 39

(23, 31, 17) (9, 11, 8)

FAP221 A8J6X7 7, 9, 7 0, 0, 0 0.0 0.015 C1d 10,28

(Pcdp1) (6, 5, 12) (0, 0, 0)

FAP114 Q45QX5 7, 7, 5 0, 0, 0 0.0 0.0081 C1a/e 34

(C1a-32) (13, 8, 16) (0, 0, 0)

FAP119 Q45QX4 7, 7, 3 0, 0, 1 5.1 0.0034 C1a/e 34

(C1a-34) (8, 7, 5) (0, 0, 1)

FAP297 A8HQE0 4, 8, 2 0, 0, 0 0.0 0.0040 C1d 28

(WDR93) (6, 6, 3) (0, 0, 0)

PF20 A8ITB4 3, 7, 2 1, 0, 0 9.9 0.028 C1-C2 bridge 44

(3, 6, 3) (1, 0, 0)

PP1a† Q9XGU3 2, 2, 1 0, 0, 0 0.0 < 0.00010 C1 48

(2, 2, 2) (0, 0, 0)

FAP174† A8I439 1, 2, 4 3, 2, 4 79 0.75 C2 45

(2, 4, 13) (5, 2, 8) C1b/f 20

Calmodulin† A8IDP6 1, 3, 3 4, 4, 4 160 0.34 C1a/e 33,34

(1, 2, 7) (6, 5, 5)

FAP227 Q45QX6 n.d. n.d. - - C1a/e 34

(C1a-18)

Three values from biological triplicate for both exclusive unique peptide counts and quantitative values are shown.

Daggers denote that these proteins are shared with other compartments of the axoneme.

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Table 2.7.2: MS results of new CP proteins and their localizations inside the CP complex.

pf15/WT ratio WT pf15 (%) p-values Uniprot Names exclusive unique exclusive unique Localizations ID (quantitative (WT vs peptide count peptide count values were pf15) (quantitative value) (quantitative value) used)

CHLREDRAFT_150638 A8J566 1, 5, 0 (1, 4, 0) 0, 0, 0 (0, 0, 0) 0 0.23 C1a/e (this study)

CHLREDRAFT_170023 A8IMQ8 3, 9, 0 (3, 6, 0) 0, 0, 0 (0, 0, 0) 0 0.17 C1b/f (this study)

CHLREDRAFT_177061 A8J9A4 7, 7, 2 (6, 5 ,3) 0, 0, 1 (0, 0, 1) 7.1 0.0098 C1b/f (this study)

C1a/e DPY30** A8J1X7 1, 2, 1 (1, 1, 2) 0, 0, 0 (0, 0, 0) 0.0 0.0041 (20 & this study)

EF-3 A8ISZ1 4, 2, 1 (4, 1, 2) 0, 0, 0 (0, 0, 0) 0.0 0.047 Not assigned

FAP7* A8IVW2 14, 16, 7 (26, 33, 28) 1, 0, 1 (1, 0, 1) 2.6 0.00025 C1a/e (this study)

FAP47** A8IPW8 22, 35, 15 (23, 27, 26) 0, 0, 0 (0, 0, 0) 0.0 < 0.00010 C2b (this study)

C2a,c,d,e and Bridge FAP65* A8JFU2 13, 23, 10 (12, 19, 16) 0, 0, 0 (0, 0, 0) 0.0 0.0016 (this study)

C2a,c,d,e and Bridge FAP70* A8I7W0 12, 22, 11 (14, 26, 23) 0, 1, 0 (0, 1, 0) 33 0.0053 (this study)

C2a,c,d,e and Bridge FAP75* A8HYW3 13, 19, 8 (13, 16, 15) 0, 1, 1 (0, 1, 1) 5.0 < 0.00010 (this study)

C1a-e-c complex FAP76** A8J128 24, 26, 14 (26, 23, 26) 0 ,1, 0 (0, 1, 0) 1.5 < 0.00010 (42 and this study)

C1a-e-c complex FAP81* A8IPC1 23, 27, 12 (24, 24, 23) 0, 0, 0 (0, 0, 0) 0.0 < 0.00010 (42 and this study)

C1a-e-c complex FAP92* A8HR45 28, 30, 17 (29, 23, 36) 0, 0, 0 (0, 0, 0) 0.0 0.0017 (42 and this study)

FAP99** A8IUG5 9, 13, 1 (10, 9, 2) 0, 0, 0 (0, 0, 0) 0.0 0.060 C1d (this study)

C1a-e-c complex FAP105* A8IKV8 3, 5, 0 (3, 4, 0) 0, 0, 0 (0, 0, 0) 0.0 0.12 (42 and this study)

C1a-e-c complex FAP108* A8IPA9 2, 3, 1 (2, 2, 2) 0, 0, 0 (0, 0, 0) 0.0 0.00090 (42 and this study)

C1a-e-c complex FAP123* A8IEJ6 4, 3, 0 (4, 2, 0) 0, 0, 0 (0, 0, 0) 0.0 0.16 (this study)

FAP125* A8IY87 14, 18, 9 (15, 18, 16) 0, 0, 0 (0, 0, 0) 0.0 < 0.00010 C1c (this study)

C2a,c,d,e and Bridge FAP147* A8IT32 7, 10, 3 (6, 7, 7) 0, 0, 0 (0, 0, 0) 0.0 < 0.00010 (this study)

C2a,c,d,e and Bridge FAP171* A8IUF4 4, 9, 2 (4 ,7, 3) 1, 0, 0 (1, 0, 0) 8.4 0.029 (this study)

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FAP173 A8JAF7 3, 3, 1 (4, 2, 2) 0, 0, 0 (0, 0, 0) 0.0 0.012 Not assigned

C2a,c,d,e and Bridge FAP194* A8J5U4 9, 13, 4 (10, 10, 7) 0, 0, 0 (0, 0, 0) 0.0 0.0013 (this study)

FAP199 A8J1E6 1, 2, 3 (1, 1, 7) 0, 0, 0 (0, 0, 0) 0.0 0.19 Not assigned

FAP209 A8J100 6, 8, 3 (6, 5, 5) 0, 1, 0 (0,1,0) 6.7 0.00086 C1c (this study)

C1a-e-c complex 42 FAP216* A8JGM3 12 ,16, 4 (13, 13, 7) 0, 0, 0 (0, 0, 0) 0.0 0.0064 C1a/e (this study)

FAP219 A8J9I0 5, 7, 1 (5, 5, 2) 0, 0, 0 (0, 0, 0) 0.0 0.022 C1c (this study)

C2a,c,d,e and Bridge FAP225* A8HNF2 14, 19, 4 (14, 22, 7) 0, 0, 0 (0, 0, 0) 0.0 0.034 (this study)

C2a,c,d,e and Bridge FAP239* A8J319 0, 5, 2 (0, 3, 3) 0, 0, 0 (0, 0, 0) 0.0 0.12 (this study)

tether complex of I1 dynein of IDA 40,41

FAP244† A8IZG0 12, 14, 5 (14, 10, 12) 0, 0, 0 (0, 0, 0) 0.0 0.00069 C2a,c,d,e and Bridge

(this study)

C1b/f FAP246** A8HNZ7 7, 6, 3 (7, 6, 5) 0, 0, 0 (0, 0, 0) 0.0 0.00096 (20 & this study)

C2a,c,d,e and Bridge FAP266* A8JB69 4, 5, 2 (4, 3 ,3) 1, 1, 0 (1, 1, 0) 23 0.0043 (this study)

C1a-e-c complex FAP279 A8HWC6 5, 7, 1 (6, 4, 2) 0, 0, 0 (0, 0, 0) 0.0 0.029 (this study)

C1a-e-c complex FAP289* A8JCZ9 8, 12, 4 (9, 9, 8) 0, 0, 0 (0, 0, 0) 0.0 < 0.00010 (this study)

FAP312* A8IUV6 2, 5, 1 (2, 3, 2) 0, 0, 0 (0, 0, 0) 0.0 0.0066 C2 20

FAP348* A8JBI2 2, 3, 2 (2, 2, 3) 0, 0, 0 (0, 0, 0) 0.0 0.0095 C1a/e (this study)

FAP412* A8JGL8 6, 9, 0 (6, 6, 0) 0, 0, 0 (0, 0, 0) 0.0 0.12 C1a-e-c complex 42

MOT17* A8J798 3, 4, 3 (3, 2, 7) 0, 0, 0 (0, 0, 0) 0.0 0.044 C1c (this study)

Membrane + matrix fraction [38] Phosphoglycerate A8HVU5 4, 10, 0 (4, 7, 0) 0, 0, 0 (0, 0, 0) 0.0 0.15 mutase†

C1b/f? (this study)

*: CP candidates shared with 20. **: CP proteins which were confirmed by 20. †: CP proteins shared with other axonemal compartments. Three values from biological triplicate for both exclusive unique peptide counts and quantitative values are shown. Proteins that has 2 or more of mean values of detected exclusive unique peptides in WT are listed, except DPY30 which was confirmed as a CP protein in 20.

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2.8 Figures

Figures 2.8.1: Structures of the axoneme and the CP complex.

(A and B) Schematic diagrams of the axoneme (A) and the CP complex structure (B) viewed from the flagellar base. The axoneme structure consists of nine DMTs radially surrounding the CP complex. The DMTs are decorated with ODA, IDA and RS complexes. IFT trains are transported at the space between the membrane and the DMTs. The CP consists of two structurally dimorphic singlets termed C1 and C2 and are connected by the bridge. Several distinct sub-structures bind around the singlets with a repeating pattern along the axis of the axoneme. Diagonal link is also known to connect the C2 with the C1b region. The model of the CP structure is adopted from 15.

(C) Cryo-ET of purified WT Chlamydomonas axoneme. Cross section (left) and longitudinal

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section (right) of 3D volume of demembranated WT axoneme are shown. Our purified WT axoneme retained both singlet microtubules of the CP complex with protruding sub-structures after demembranation step as shown in red arrowheads. Yellow dashed line indicates the section in the right panel. Scale bar represents 50 nm.

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Figure 2.8.2: Preparation of microtubule fraction for MS.

(A) A schematic diagram of sequential purification of the axoneme. Flagella were demembranated using the detergent NP-40 following the isolation from Chlamydomonas cells. Demembranated axonemes were incubated with ADP and ATP to induce splitting of the DMTs and the CP. The samples were then treated with 0.6 M NaCl twice to shed large protein complexes such as dyneins.

Note that illustrations here show protein compositions rather than the actual structures. (B) SDS-

PAGE gel demonstrating protein shedding after sequential purification. The signal of the dynein heavy chain band (> 500 kDa) was decreased significantly after NaCl treatments. In contrast, the tubulin band which is a main component of the CP and DMTs showed little change after sequential purification. (C) A typical cryo-EM image of purified sample showing the presence of singlet microtubule from the CP. In our cryo-electron micrographs of our

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purified microtubule fraction, both DMTs (orange arrowheads) and singlet microtubule from the

CP (red arrowhead) with characteristic protruding sub-structures were observed (see also Fig.

S2.9.2). Boxed area of the micrograph is shown in the right panel. The plot profile of yellow box area was generated by ImageJ and the distances between the peaks (red dots) were measured. The averaged distance between the protrusions was 16.7 nm which is consistent with the known repeating unit of the CP 15. There were more numbers of DMTs compared with singlets from the

CP reflecting the stoichiometry inside the axoneme. Scale bar represents 100 nm.

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Figure 2.8.3: Identification of new CP proteins by MS.

(A) Schematic diagrams of the axoneme structures from WT and pf15 Chlamydomonas strains and expected protein compositions obtained from either axoneme structure. The DMTs and the CP complex proteins would be purified from WT flagella while only the proteins from DMTs are expected to be purified from pf15 flagella. Obtained microtubule fractions from WT and pf15 were analyzed by MS and the results were compared. (B) SDS-PAGE result of sequential purification of microtubule fraction from pf15 flagella showing similar pattern with that of WT flagella. (C) A volcano plot comparing WT and pf15 MS results. Changes in a protein abundance between WT (n

= 3) and pf15 (n = 3) results were plotted. A dashed red line indicates the significance threshold of p < 0.05 and proteins meet this criterion are shown in green. Triangle dots represent completely missing proteins in either WT or pf15 result. Two- and four-fold changes are shown by the orange and blue dashed lines, respectively. There were more proteins completely missing in pf15 results while many others showed more than two-fold decrease in pf15 results. (D) Plot of fold changes of proteins categorized into different groups. Proteins identified by MS were arranged in groups (Tubulins; RS proteins; IFT complex proteins; IFT dynein; IFT kinesin; axonemal dyneins; dynein regulatory complex; MIP candidates and known CP proteins) and fold change between WT and pf15 results of each protein was plotted. Two- and four-fold changes are shown by the orange and blue dashed lines, respectively. Green lines indicate the median value for each category. Statistical significance compared with tubulin result was examined by one-way ANOVA followed by Dunnett’s multiple comparisons test. Among these classes, only known CP protein class were significantly reduced with a p-value of 0.00050. Fold changes of our CP protein candidates are also shown at the rightmost column. Red line represents proteins that were

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completely missing in pf15. Proteins included in each class are listed in Tables 2.7.1, 2.7.2 and

S2.9.4.

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Figure 2.8.4: MS analyses using Chlamydomonas mutant strains lacking CP sub-structures.

(A) Schematic diagrams of CP compositions from mutants lacking sub-structures of the CP.

Sub-structures of the CP which are missing in pf6, cpc1 and pf16 strains are shown in dashed lines. pf6 strain is missing the C1a/e structure (formerly the C1a), cpc1 strain lacks the C1b/f structure

(formerly the C1b) while pf16 strain has an unstable C1 structure. In the pf16 model, The C1b/f region is in transparent since this region can remain attached to the C2 microtubule with the

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diagonal link 14. Note that these diagrams show the protein compositions in the samples rather than the actual structures. (B-I) MS profiles of CP proteins and their possible localizations. Detected levels of proteins were compared among strains (WT, pf6, cpc1, pf16 and pf15). Mean values of normalized quantitative values of each CP protein are shown (error bars represent SD for biological triplicate). Known CP proteins that have been localized to specific sub- complexes showed similar MS profiles. These proteins were used as references to assign newer

CP candidates to certain sub-structures, such as the C1a/e area (B), the C1c area (C), the C1a-e-c complex (D), the C1d region (E), the C1b/f area (F), the C2b area (G), the C2a, c, d, e & bridge area (H), and proteins shared with other axonemal structures (I). Known CP proteins are labelled in black, CP candidates shared with 20 are labelled in blue, and CP candidates obtained only in our work are in red. One-way ANOVA followed by Dunnett’s multiple comparisons test comparing with WT values (*p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; n.s., not significant; n.d., not detected). Plots not shown here are presented in Fig. S2.9.4.

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Figure 2.8.5: Model of localizations of CP proteins.

Proteins are mapped to CP sub-structures based on our MS profiles. Traditionally known CP proteins are shown in black, CP candidates shared with 20 are in blue and proteins identified only in our results are in red. Daggers denote the proteins possibly shared with other axonemal structures.

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2.9 Supplementary Material

Supplementary Fig. 2.9.1: Cross-sectional EM results.

(A) Cross-sectional EM images of untreated WT flagella. Axonemes with two singlet microtubules from the CP complex were observed along with axonemes with only one of the CP microtubules, presumably the C1 microtubule. (B) EM images of cross-section of WT sample after demembranation, ADP/ATP treatment and twice NaCl treatment. The majority of the axoneme structure had only one of the CP microtubules, which is thought to correspond to the C1 |70|

microtubule. Inset in the left panel shows the magnified view in the right panel. (C) Cross-sectional

EM images of untreated pf16 flagella. The majority of the axonemes had only one of the CP microtubules, which is thought to be the C2 microtubule. (D) EM images of cross-section of pf16 sample after demembranation, ADP/ATP treatment and NaCl treatments. Most of the axonemes had no clear microtubule structures from the CP but the residual materials were observed inside the axonemes. Note that there are more split DMTs and the CP microtubules after ADP/ATP treatment. (E) Quantification of the numbers of microtubules of the CP observed in the axonemes.

The numbers of axonemes used for the quantification were as follows: untreated WT, n=668; treated WT, n=1,322; untreated pf16, n=323; treated pf16, n=878.

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Supplementary Fig. 2.9.2: Additional cryo-EM images showing singlets from the CP.

In our purified microtubule fractions, we occasionally observed singlet microtubules from the CP

(red arrowheads) with characteristic appendages along with the DMTs (orange arrowheads). Scale bars, 100 nm.

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Supplementary Fig. 2.9.3: Purification of microtubule fractions from cpc1, pf6 and pf16 strains, and comparison of MS results with WT.

(A and B) SDS-PAGE results of sequential purifications of microtubule fractions from cpc1 flagella and a volcano plot comparing cpc1 result with WT result. (C and D) SDS-PAGE gel of

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purification from pf6 axoneme and a volcano plot of pf6 MS result compared with WT result. (E and F) pf16 strain SDS-PAGE result and its MS result on a volcano plot compared with WT.

Dashed red line indicates the significance threshold (p < 0.05) and proteins showed significant change are shown in green. Triangle dots represent completely missing proteins in either mutant results or WT result. Orange and blue dashed lines indicate two-and four-fold changes, respectively.

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Supplementary Fig. 2.9.4: Additional MS profiles for other CP protein candidates.

MS profiles of CP candidates which were not readily assigned to certain CP sub-structures are summarized here.

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Supplementary Table 2.9.1: Proteins increased in pf15 compared with WT.

WT pf15

p-values exclusive unique exclusive unique Names Uniprot ID Comments peptide count peptide count (WT vs pf15)

(quantitative value) (quantitative value)

Proteins identified by at least a 2-4 fold increase

CHLREDRAFT_144597 A8IK96 12, 13, 2 15, 15, 14 0.0081

(13, 11, 5) (25, 27, 21)

CHLREDRAFT_144760 A8ILI2 4, 12, 3 9, 12, 9 0.021

(4, 7, 5) (11, 14, 9)

CHLREDRAFT_173135 A8IUI7 1, 3, 1 3, 4, 4 0.0024

(1, 2, 2) (4, 5, 4)

FAP143 A8JEG3 3, 5, 3 6, 5, 8 0.027 Microtubule Inner

Protein [1] (4, 6, 7) (13, 9, 16)

FAP200 (DRC8) A8J3A0 2, 2, 0 2, 2, 3 0.040 DRC protein

(3, 2, 0) (5, 3, 5) [2]

FAP207 A8IHH6 7, 5, 1 9, 10, 8 0.036 Axoneme

(8, 4, 2) (11, 11, 10) [3]

FAP272 A8IDN4 1, 4, 0 4, 3, 3 0.045

(1, 3, 0) (5, 3, 4)

FBB9 A8J795 3, 3, 1 5, 4, 6 0.00079 Flagellar/basal

body protein (4, 2, 2) (8, 8, 8)

IFT22 (FAP9) A8HME3 3, 5, 0 5, 4, 5 0.021 IFT

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(3, 4, 0) (7, 6, 8)

IFT56 (DYF13) A8JA42 5, 8, 2 8, 9, 8 < 0.00010 IFT

(6, 5, 5) (11, 11, 11)

IFT57 Q2XQY7 1, 2, 0 2, 2, 2 0.014 IFT

(1, 1, 0) (2, 2, 3)

IFT74 Q6RCE1 6, 16, 3 14, 15, 19 0.021 IFT

(6, 14, 7) (20, 18, 26)

RSP16 A8IKR9 17, 15, 16 18, 17, 23 0.00045 RSP protein

(27, 21, 20) (50, 50, 55)

Proteins identified by at least a 4 fold increase

BBS1 A8JEA1 0, 1, 0 1, 4, 2 0.049 BBSome protein

(0, 1, 0) (1, 5, 3)

CHLREDRAFT_168407 A8HRH6 1, 0, 0 2, 2, 2 0.0038

(1, 0, 0) (2, 2, 2)

CNK8 A8IHF8 2, 2, 0 6, 4, 3 0.030 NimA-related

protein kinase 8 (2, 1, 0) (7, 6, 3)

FAP260 A8IR95 3, 3, 3 18, 23, 16 0.0035 Flagellar

associated (4, 2, 5) (27, 34, 21) membrane protein

RPP0 A8J5Z0 0, 1, 0 1, 2, 3 0.028 60S acidic

ribosomal protein (0, 1, 0) (1, 2, 3) P0

RPS8 A8HVQ1 0, 1, 0 2, 2, 3 0.0020 40S ribosomal

protein S8 (0, 1, 0) (2, 2, 3)

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Proteins only detected in pf15

CHLREDRAFT_141055 A8ICE6 0, 0, 0 1, 3, 3 0.021

(0, 0, 0) (1, 3, 3)

CHLREDRAFT_142741 A8I6P0 0, 0, 0 2, 2, 2 < 0.00010

(0, 0, 0) (2, 2, 2)

CHLREDRAFT_142766 A8I6V5 0, 0, 0 4, 2, 1 0.071

(0, 0, 0) (5, 2, 1)

CHLREDRAFT_153169 A8JDK3 0, 0, 0 4, 7, 6 0.0022

(0, 0, 0) (5, 8, 7)

CHLREDRAFT_167270 A8HX54 0, 0, 0 2, 2, 2 0.00064

(0, 0, 0) (2, 2, 3)

CHLREDRAFT_168271 A8I1D6 0, 0, 0 3, 4, 1 0.043

(0, 0, 0) (4, 5, 1)

CHLREDRAFT_179827 A8JG61 0, 0, 0 4, 4, 0 0.12

(0, 0, 0) (5, 5, 0)

Values from biological triplicate are shown.

Proteins whose mean values of detected exclusive unique peptides are 2 or more in pf15 result are listed here.

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Supplementary Table 2.9.2: List of proteins only identified in Zhao et al., (2019). [4]

WT pf15

Names Uniprot ID exclusive unique peptide exclusive unique peptide Comments

count (quantitative value) count (quantitative value)

FAP39 A8J0V2 6, 10, 4 (8 ,9 ,7) 4, 5, 4 (8 ,10, 4) Not significantly

decreased in pf15

FAP49 A8J4C7 11, 0, 0 (10, 0, 0) 9, 8, 9 (11, 9, 9) Not confident*

FAP72 A8J4C5 10, 3, 0 (9, 2, 0) 9, 8, 9 (11, 9, 9) Not confident*

FAP139 A8J134 11, 16, 2 (12, 15 ,5) 8, 9 ,8 (10, 16, 8) Not significantly

decreased in pf15

FAP154 A8J4C9 18, 4, 4 (17, 2, 7) 16, 14, 19 (19, 16, 20) Not confident*

FAP178 A8ID60 0, 2, 0 (0, 2, 0) 0 ,0, 0 (0, 0, 0) Not significantly

decreased in pf15

FAP275 A8J870 0, 1, 0 (0, 1, 0) 0, 0, 0 (0, 0, 0) Not significantly

decreased in pf15

FAP286 A8IKJ2 0 ,0, 0 (0, 0, 0) 0 ,0, 0 (0, 0, 0) Not detected

FAP345 A8JED5 1, 1, 2 (1, 1, 3) 0, 0, 0 (0, 0, 0) Not significantly

decreased in pf15

FAP380 A8HP72 2, 1, 0 (2, 1, 0) 1, 0, 0 (1, 0, 0) Not significantly

decreased in pf15

FAP411 A8J4L4 0 ,0, 0 (0, 0, 0) 0 ,0, 0 (0, 0, 0) Not detected

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FAP413 A8J3X1 0, 0, 2 (0, 0, 5) 0, 0, 0 (0, 0, 0) Not significantly

decreased in pf15

FAP414 A8J0A0 0, 2, 0 (0, 1, 0) 1, 0, 0 (1, 0, 0) Not significantly

decreased in pf15

FAP415 A8I0B9 0 ,0, 0 (0, 0, 0) 0 ,0, 0 (0, 0, 0) Not detected

FAP416 A8IQU8 0 ,0, 0 (0, 0, 0) 0 ,0, 0 (0, 0, 0) Not detected

FAP417 A8IBK1 1, 0, 0 (1, 0, 0) 0, 1, 0 (0, 1, 0) Not significantly

decreased in pf15

DIP13 Q9XF62 0 ,0, 0 (0, 0, 0) 0 ,0, 0 (0, 0, 0) Not detected

NAP O24426 0 ,0, 0 (0, 0, 0) 0 ,0, 0 (0, 0, 0) Not detected

Three values from biological triplicate for both exclusive unique peptide counts and quantitative values are shown.

*: There were peptide sequences shared between FAP49, FAP72 and FAP154 and the detected peptides were not confidently assigned to each protein. Therefore, total spectrum counts are shown for these proteins instead of exclusive unique peptide counts.

1. Ma, M., et al., Structure of the Decorated Ciliary Doublet Microtubule. Cell, 2019. 179(4):

p. 909-922 e12.

2. Gui, L., et al., Scaffold subunits support associated subunit assembly in the

Chlamydomonas ciliary nexin-dynein regulatory complex. Proc Natl Acad Sci U S A,

2019. 116(46): p. 23152-23162.

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3. Lechtreck, K.F., et al., HA-tagging of putative flagellar proteins in Chlamydomonas

reinhardtii identifies a novel protein of intraflagellar transport complex B. Cell Motil

Cytoskeleton, 2009. 66(8): p. 469-82.

4. Zhao, L., et al., Proteome of the central apparatus of a ciliary axoneme. J Cell Biol, 2019.

218(6): p. 2051-2070.

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17 Lechtreck, K. F., Delmotte, P., Robinson, M. L., Sanderson, M. J. & Witman, G. B. Mutations in Hydin impair ciliary motility in mice. J Cell Biol 180, 633-643, doi:10.1083/jcb.200710162 (2008). 18 Li, X. et al. An Indexed, Mapped Mutant Library Enables Reverse Genetics Studies of Biological Processes in Chlamydomonas reinhardtii. The Plant cell 28, 367-387, doi:10.1105/tpc.15.00465 (2016). 19 Teves, M. E., Nagarkatti-Gude, D. R., Zhang, Z. & Strauss, J. F., 3rd. Mammalian axoneme central pair complex proteins: Broader roles revealed by gene knockout phenotypes. Cytoskeleton (Hoboken) 73, 3-22, doi:10.1002/cm.21271 (2016). 20 Zhao, L., Hou, Y., Picariello, T., Craige, B. & Witman, G. B. Proteome of the central apparatus of a ciliary axoneme. J Cell Biol 218, 2051-2070, doi:10.1083/jcb.201902017 (2019). 21 Dymek, E. E., Lefebvre, P. A. & Smith, E. F. PF15p is the chlamydomonas homologue of the Katanin p80 subunit and is required for assembly of flagellar central microtubules. Eukaryot Cell 3, 870-879, doi:10.1128/EC.3.4.870-879.2004 (2004). 22 Dutcher, S. K., Huang, B. & Luck, D. J. Genetic dissection of the central pair microtubules of the flagella of Chlamydomonas reinhardtii. J Cell Biol 98, 229-236, doi:10.1083/jcb.98.1.229 (1984). 23 Smith, E. F. & Lefebvre, P. A. PF16 encodes a protein with armadillo repeats and localizes to a single microtubule of the central apparatus in Chlamydomonas flagella. J Cell Biol 132, 359-370 (1996). 24 Goduti, D. J. & Smith, E. F. Analyses of functional domains within the PF6 protein of the central apparatus reveal a role for PF6 sub-complex members in regulating flagellar beat frequency. Cytoskeleton (Hoboken) 69, 179-194, doi:10.1002/cm.21010 (2012). 25 Gorman, D. S. & Levine, R. P. Cytochrome f and plastocyanin: their sequence in the photosynthetic electron transport chain of Chlamydomonas reinhardi. Proc Natl Acad Sci U S A 54, 1665-1669, doi:10.1073/pnas.54.6.1665 (1965). 26 Craige, B., Brown, J. M. & Witman, G. B. Isolation of Chlamydomonas flagella. Current protocols in cell biology Chapter 3, Unit-3.41.49, doi:10.1002/0471143030.cb0341s59 (2013). 27 Shevchenko, A., Tomas, H., Havli, J., Olsen, J. V. & Mann, M. In-gel digestion for mass spectrometric characterization of proteins and proteomes. Nature Protocols 1, 2856, doi:10.1038/nprot.2006.468 (2007). 28 Brown, J. M., Dipetrillo, C. G., Smith, E. F. & Witman, G. B. A FAP46 mutant provides new insights into the function and assembly of the C1d complex of the ciliary central apparatus. J Cell Sci 125, 3904-3913, doi:10.1242/jcs.107151 (2012). 29 Johnson, K. A., Haas, M. A. & Rosenbaum, J. L. Localization of a kinesin-related protein to the central pair apparatus of the Chlamydomonas reinhardtii flagellum. J Cell Sci 107 ( Pt 6), 1551-1556 (1994). 30 Lechtreck, K. F. & Witman, G. B. Chlamydomonas reinhardtii hydin is a central pair protein required for flagellar motility. J Cell Biol 176, 473-482, doi:10.1083/jcb.200611115 (2007). 31 Pazour, G. J., Agrin, N., Leszyk, J. & Witman, G. B. Proteomic analysis of a eukaryotic cilium. The Journal of Cell Biology 170, 103-113, doi:10.1083/jcb.200504008 (2005).

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32 Ichikawa, M. et al. Subnanometre-resolution structure of the doublet microtubule reveals new classes of microtubule-associated proteins. Nat Commun 8, 15035-15035, doi:10.1038/ncomms15035 (2017). 33 Yang, P. et al. Radial spoke proteins of Chlamydomonas flagella. J Cell Sci 119, 1165- 1174, doi:10.1242/jcs.02811 (2006). 34 Wargo, M. J., Dymek, E. E. & Smith, E. F. Calmodulin and PF6 are components of a complex that localizes to the C1 microtubule of the flagellar central apparatus. J Cell Sci 118, 4655-4665, doi:10.1242/jcs.02585 (2005). 35 Ichikawa, M. et al. Tubulin lattice in cilia is in a stressed form regulated by microtubule inner proteins. Proc Natl Acad Sci U S A 116, 19930-19938, doi:10.1073/pnas.1911119116 (2019). 36 Ma, M. et al. Structure of the Decorated Ciliary Doublet Microtubule. Cell 179, 909-922 e912, doi:10.1016/j.cell.2019.09.030 (2019). 37 Lechtreck, K. F., Gould, T. J. & Witman, G. B. Flagellar central pair assembly in Chlamydomonas reinhardtii. Cilia 2, 15, doi:10.1186/2046-2530-2-15 (2013). 38 Cole, D. G. in The Chlamydomonas Sourcebook (Second Edition) (eds Elizabeth H. Harris, David B. Stern, & George B. Witman) 71-113 (Academic Press, 2009). 39 Mitchell, B. F., Pedersen, L. B., Feely, M., Rosenbaum, J. L. & Mitchell, D. R. ATP production in Chlamydomonas reinhardtii flagella by glycolytic enzymes. Mol Biol Cell 16, 4509-4518, doi:10.1091/mbc.e05-04-0347 (2005). 40 Fu, G. et al. The I1 dynein-associated tether and tether head complex is a conserved regulator of ciliary motility. Mol Biol Cell 29, 1048-1059, doi:10.1091/mbc.E18-02-0142 (2018). 41 Kubo, T., Hou, Y., Cochran, D. A., Witman, G. B. & Oda, T. A microtubule-dynein tethering complex regulates the axonemal inner dynein f (I1). Mol Biol Cell 29, 1060-1074, doi:10.1091/mbc.E17-11-0689 (2018). 42 Fu, G. et al. Structural organization of the C1a-e-c supercomplex within the ciliary central apparatus. J Cell Biol 218, 4236-4251, doi:10.1083/jcb.201906006 (2019). 43 Bloch, M. A. & Johnson, K. A. Identification of a molecular chaperone in the eukaryotic flagellum and its localization to the site of microtubule assembly. J Cell Sci 108 ( Pt 11), 3541-3545 (1995). 44 Smith, E. F. & Lefebvre, P. A. PF20 gene product contains WD repeats and localizes to the intermicrotubule bridges in Chlamydomonas flagella. Molecular biology of the cell 8, 455-467 (1997). 45 Rao, V. G. et al. Myc-binding protein orthologue interacts with AKAP240 in the central pair apparatus of the Chlamydomonas flagella. BMC Cell Biol 17, 24, doi:10.1186/s12860- 016-0103-y (2016). 46 Rupp, G., O'Toole, E. & Porter, M. E. The Chlamydomonas PF6 locus encodes a large alanine/proline-rich polypeptide that is required for assembly of a central pair projection and regulates flagellar motility. Mol Biol Cell 12, 739-751, doi:10.1091/mbc.12.3.739 (2001). 47 Bernstein, M., Beech, P. L., Katz, S. G. & Rosenbaum, J. L. A new kinesin-like protein (Klp1) localized to a single microtubule of the Chlamydomonas flagellum. J Cell Biol 125, 1313-1326, doi:10.1083/jcb.125.6.1313 (1994).

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48 Yang, P., Fox, L., Colbran, R. J. & Sale, W. S. Protein phosphatases PP1 and PP2A are located in distinct positions in the Chlamydomonas flagellar axoneme. J Cell Sci 113 ( Pt 1), 91-102 (2000).

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CHAPTER 3 - General discussion, summary, and conclusion

3.1 General discussion and summary

At the start of my degree only 22 proteins out of hundreds of axonemal proteins identified through MS have been found to localize to the CP20,75. Towards the end of my degree two separate studies including my own were able to identify over 40 additional CP proteins with 26 shared between the two studies, thus, increasing the total amount of CP proteins over three-fold75. In the second chapter, in addition to identifying new CP proteins I was able to localize these proteins around the CP as well. In conjunction with the study done by Zhao et al (2019)., and their functional assay as well as their MS study a more comprehensive understanding of the CP architecture has been achieved.

With our increased understanding of the CP, several long-standing questions may finally be addressed. The organization of CP protein complexes and their interaction with neighboring complexes has always been ambiguous. The first high resolution study of the CP by cryo-electron tomography by Carbajal-González et al (2013)., revealed for the first time the many different protein complexes and their unique repeating periodicities in Chlamydomonas reinhardtii26. It was from this structure that the naming and distinction of CP complexes were derived from. The overall resolution of the CP estimated by the Fourier Shell correlation method was estimated to be around

33Å (3.3nm)76. This resolution, however, was not good enough to go beyond describing the overall shape and pattern of the CP. In fact, the coloration and distinction of the CP and its complexes which have become the conventional way of describing the CP is largely based off arbitrary coloration. At 33Å resolution a great deal of things cannot be observed including the borders between different protein complexes. It was, therefore, the task of the author to implement his own borders based on the available data. In this case it is unsure if there is an over or underestimation

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of the protein complexes. Rather than 11 smaller protein complexes binding around the CP we have evidence that a larger super-protein complex may be at play. It was proposed in a previous study that when using PF16 as a protein bait a super complex of CP proteins was formed corresponding to the C1a, e and c region of the CP77. The protein proposed to be a part of this complex included flagellar associated protein 76, 81 92 and 21677. This finding was consistent with our MS based localization of newly identified CP proteins. The majority of protein held a MS profile suggesting it belonged to the C1a/e region where it was proposed to form this super complex. In our data there is further evidence to support the possibility of other super complexes however, more biochemical experiments need to be conducted in order to prove these conclusions.

One interesting CP protein identified by both groups was FAP125. Previously two proteins were shown to be associated to the CP. One of these proteins termed C1 kinesin had a mass of approximately 110KD (determined by SDS-PAGE) and was recognized by two different anti- peptide polyclonal antibodies against a conserved kinesin sequence78. Up until our study the association of an additional kinesin motor protein alongside KLP1 was largely assumed and never proven directly. In both our study and Zhao et al (2019)., the peptide sequence of FAP125 was identified in WT MS and was significantly reduced in the CP-less mutant pf15 in ours. We were able to identify FAP125 and exclusive to our study we were able to map its location based on the created MS profile to the region around the C1c. A proposed localization at the C1c has several particularly interesting implications. KLP1 a known kinesin motor is already known to localize at the C2c of the C2 microtubule71. FAP125 being present at the C1c would place the secondary kinesin at the mirror opposite end of the CP. The presence of an additional kinesin motor may explain why previous RNAi knock down experiment of KLP1 resulted in only reduced swim speed instead of full flagellar paralysis71. In this case it is possible that KLP1 and FAP125 serve as

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redundant motors to one another. In the future knockdowns of both KLP1 and FAP125 should be tested in order to see if flagellar paralysis can occur. The symmetrical placement of the kinesin motors may also help to regulate the motor function of dynein as previously discussed in chapter

1. Symmetrical distribution of kinesin in addition to possible rotational based regulation of RS and dynein motor protein may play an important role in how signals for movement are propagated in the CP.

With a more comprehensive understanding of the CP proteome the conserved proteins present in human have been easier to identify. Surprisingly, there is both a population of conserved proteins and species-specific proteins at the CP. This is consistent with different forms of movement unique to different species perhaps requiring a different set of regulatory proteins.

Among the newly identified conserved proteins FAP81 and FAP7 are included, both of which have been localized by us to the C1a/e region as well as potentially form a super complex between C1a- e-c complexes77. Whereas the protein FAP289 which was found to be localized near the C1e region is a species-specific variant to Chlamydomonas reinhardtii. The logical next step should be characterizing the newly identified CP proteins with human orthologs. In this way we are able to tie a functional or morphological phenotype to mutant forms of the protein. This may prove to help the rate of diagnosis in patients during a whole exome-sequencing.

3.2 Conclusion:

As awareness of ciliopathies such as PCD grows the greater the necessity it is to diagnose and treat it. The greatest challenge facing patients, physicians and researchers is the heterogeneous nature of ciliopathies like PCD. With potentially hundreds of causative genes, patients lack well-defined treatments while physicians struggle to improve diagnostic measures.

The burden of this then lies on the researcher to improve our base understanding of the mechanisms

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and proteins involved. This thesis investigates the protein composition of CP a critical structure present in motile cilia through proteomics. In doing so we were able to increase the speed at which

CP proteins were identified in order to expedite functional experiments. We have identified over

40 new CP proteins whose location within the axoneme was previously unknown. With this, we hope to have improved our overall understanding of the axoneme while hopefully making an impact at the clinical level as well.

3.3 Future plan

Moving forward the first aim should be to proceed towards the functional analysis of newly identified CP proteins. We can first obtain Chlamydomonas mutants of newly identified CP proteins from the Chlamydomonas library. From then on, we need to genetically backcross the mutant to remove unwanted background mutations. After which we may then move to evaluating the swimming and flagellar movement and observe the axoneme in cross-section with TEM.

Validation through a rescue experiment will help to finalize a newly characterized CP protein function and significance. A structural analysis of the CP and mutant variations through cyro- electron techniques such as single-particle analysis or tomography will not only be complementary to the previous aim but can help to improve the current resolution of the CP structure in WT

Chlamydomonas. Then we can begin to dissect the structure and function with similarly resolved structures from mutant CP strains.

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