Master thesis (Mette Lethan)

Identification and characterization of novel
Chlamydomonas flagellar tip proteins

Master Thesis by Mette Lethan

Department of Biology
University of Copenhagen
Denmark
2009

Supervisor
Lotte Bang Pedersen
F A C U L T Y O F S C I E N C E
U N I V E R S I T Y O F C O P E N H A G E N

Master thesis by Mette Lethan

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Preface
This master thesis represents the final part of my education and is based on
experimental work carried out from May 2008 to June 2009 at Department of Biology,
Section of Cell and Developmental Biology, University of Copenhagen.
First of all I wish to thank my supervisor and daily mentor associate professor, Ph.D.
Lotte Bang Pedersen, for letting me join in on a very interesting project, thereby
introducing me to the cilia and a very smart model organism called Chlamydomonas,
and also for always being ready with help and support all the way through the project. A
special thanks to Ph. D student Jacob M. Schrøder for helping with the NIH3T3
fibroblast cells and always being ready with a great humour and a good story. Special
thanks to technician Søren L. Johansen for helping in the lab, and always providing
what you seem to be missing. A great thanks to the entire cilia group which comprises a
gathering of amasing people. A special thanks to Dorte L. Egeberg, Sonja K. Brorsen
and Tue S. Jørgensen for daily inspiration and discussions. A big thanks to the entire 5.
Floor especially to the entire ”Grøn Stue”, you know who you are, for supplying all the of
non-”lab” related activities. Without you this year would not have been the same. I am
very grateful to Niovi Santama for generously providing me with Nubp1 antibodies as
well as the Anna Akhmanova group for collaboration in the search of IFT172 and EB1
binding partners. I also thank the Chlamydomonas Genetics Center for strains. Lastly I
wish to thank my family and all my friends, for understanding that time is scarce.
Parts of the results obtained in this project were presented with a poster at the Gordon
Research Conference on Cilia, Mucus and Mucociliary Interactions in february 2009 in
Lucca, Italy.
Copenhagen, august 2009
__________________________
Mette Lethan

Identification and characterization of novel Chlamydomonas flagellar tip proteins

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Table of Contents
Preface .....................................................................................................................................2
Abstract ...................................................................................................................................5
Dansk resume.......................................................................................................................6
Abbreviations .......................................................................................................................8
1. Aim of study ...................................................................................................................11
1.1. Introductory remarks............................................................................................11
1.2. Specific aims........................................................................................................11
2. Introduction ....................................................................................................................13
2.1. Cilia and flagella structure ...................................................................................13
2.2. Ciliopathies ..........................................................................................................16
2.3. Ciliogenesis and the cell cycle.............................................................................17
2.4. Intraflagellar transport..........................................................................................18
2.4.1. Anterograde IFT............................................................................................20
2.4.2. Retrograde IFT..............................................................................................22
2.4.3. Tip turnaround...............................................................................................23
2.4.4. IFT particle polypeptides...............................................................................24
2.4.4.1. IFT172 ....................................................................................................25
2.5. EB1......................................................................................................................26
2.6. Nucleotide-binding protein 1 (Nubp1)..................................................................27
2.7. Kinesins ...............................................................................................................28
2.7.1. Kinesin-2 family proteins...............................................................................30
2.7.2. Kinesin-5 family proteins...............................................................................31
2.7.3. Kinesin-14 family proteins.............................................................................31
2.8. Chlamydomonas as a model organism for ciliary functions ................................32
3. Results and discussion............................................................................................37
3.1. Introductory notes................................................................................................37
3.2. Chlamydomonas reinhardtii Nubp1 is present in the flagella ..............................38
3.3. CrNubp1 localizes to the soluble membrane plus matrix compartment ..............40
3.4. The flagellar level of CrNubp1 is unaffected by mutations affecting
assembly of the main axonemal substructures: outer dynein arms, inner
dynein arms, radial spokes and the central apparatus........................................42
3.5. CrNubp1 localizes to the basal bodies and the tip of the flagella ........................43
3.6. Mammalian Nubp1 localizes to the centrioles and the nucleus in NIH3T3
fibroblast cells......................................................................................................46
3.7. Identification of possible binding partners to CrEB1 and IFT172 C-term
in Chlamydomonas wild type (CC-124) flagella ..................................................47
3.8. Construction and purification of MBP-ARFA1A, MBP-FAP20 and
MBP-Eg5 motor domain fusion proteins..............................................................52


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3.9. MBP pull-down analysis of ARFA1A, FAP20 and Eg5 motor domain .................52
3.10. Further testing of the function of ARFA1A.........................................................56
4. Conclusions and future directions.....................................................................59
5. Materials and Methods..............................................................................................63
5.1. Eukaryotic cell cultures........................................................................................63
5.2. Preparation of flagella and cell body extracts......................................................65
5.3. PCR and cloning procedures...............................................................................66
5.4. Expression of MBP fusion proteins......................................................................69
5.5. Purification of fusion proteins on amylose resin ..................................................69
5.6. MBP pull-down assays ........................................................................................69
5.7. Protein quantification ...........................................................................................70
5.8. Sodium Dodecyl Sulphate PolyacrylAmide Gel Electrophoresis (SDS-PAGE) ...70
5.9. Western Blot analysis (WB).................................................................................71
5.10. Antibodies and affinity purification .....................................................................71
5.11. Immunofluorescence microscopy analysis (IFM) ..............................................72
5.11.1. IFM on Chlamydomonas cells.....................................................................74
5.11.2. IFM on NIH3T3 cells ...................................................................................75
6. References ......................................................................................................................77
7. Appendices.....................................................................................................................84
Appendix A: Culturing media ......................................................................................84
Appendix B: Preparation of flagella and cell body extracts ........................................85
Appendix C: cDNA sequences and multiple sequence alignments............................88
Appendix D: Vector map.............................................................................................93
Appendix E: Primers...................................................................................................94
Appendix F: Procedure for PCR .................................................................................95
Appendix G: Agarose gels..........................................................................................96
Appendix H: Transformation of DH10α E. coli cells ...................................................96
Appendix I: Protein quantification ...............................................................................97
Appendix J: Solutions for SDS-PAGE and western blotting .......................................99
Appendix K: Affinity purification of CrNubp1.............................................................100
Appendix L: IFM........................................................................................................102
Appendix M: Overview of potential binding partners of EB1/IFT172 from
Chlamydomonas flagella ..........................................................................................103

*Picture on front page from:
http://rydberg.biology.colostate.edu/Phytoremediation/2003/Boczon/chlamydomonas02.jpg


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Abstract
Cilia and flagella are microtubule (MT)-based organelles protruding from the cell surface
of most eukaryotic cell, which play important roles in motility and sensory signaling.
Lack of normal functioning cilia can a number of diseases and developmental defects
including polycystic kidneys, blindness and polydactyly. Assembly and maintenance of
cilia are mediated by intraflagellar transport (IFT) a highly conserved bidirectional MT-
based transport system. IFT transports flagellar precursors from the flagellar base to the
tip for assembly (anterograde transport) and returns turnover products from the tip back
to the base (retrograde transport). The IFT system consists of anterograde (kinesin-2)
and retrograde (cytoplasmic dynein 2) motor complexes, and ca. 17 different IFT
particle proteins separated in two large complexes, A and B. The molecular
mechanisms by which these different components are coordinated and regulated at the
flagellar base and tip are unclear.
The unicellular green alga Chlamydomonas reinhardtii is a well-established
model organism for studying cilia and IFT. IFT turnaround at the flagellar tip involves:
inactivation/down-regulation of kinesin-2, activation/upregulation of cytoplasmic dynein
2, unloading of flagellar precursors and loading of flagellar turnover products. It has
previously been shown that EB1, a small MT plus-end tracking protein (+TIP) localizes
to the flagellar tip in Chlamydomonas where it interacts with IFT172 possibly regulating
IFT particle turnover. The aim of this project was to characterize and identify flagella tip
proteins, which are presumed to play central roles in IFT regulation and/or cilia
assembly and function. To this end, I used Chlamydomonas as a model organism and I
employed two different strategies. First, using an antibody generated against the small
nucleotide-binding protein 1 (Nubp1) prior to the onset of this study, I show using
western blotting and immunofluorescence microscopy that Nubp1 is localized to the
flagella in Chlamydomonas and is specifically enriched at the flagellar tip. Second, I set
out to identify binding partners of EB1 and IFT172 C-terminus using GST pull-down of
isolated flagella. This part of my thesis work was done in collaboration with Anna
Akhmanova and her group in Rotterdam, The Netherlands. Akhmanovas group
executed the GST pull-down experiments in Chlamydomonas using isolated flagella and


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GST-EB1/IFT172 fusion proteins and identified putative binding partners by mass
spectrometry. I subsequently cloned and characterized three of the potential
EB1/IFT172 binding partners identified.
Dansk resume
Cilier og flageller er mikrotubuli (MT)-baserede organeller der udgår fra celleoverfladen
hos de fleste eukaryote celler, hvor de spiller vigtige roller i bevægelse og sensorisk
signalering. Hvis normalt fungerende cilier mangler kan det føre til forskellige
sygdomme og udviklingsmæssige defekter inklusiv cystenyre, blindhed og polydaktyli.
Dannelse og vedligeholdelse af cilier er medieret af intraflagellær transport (IFT) et
meget konserveret bi-direktionelt MT-baseret transport system. IFT transporterer de
flagellære byggesten fra flagellets base og til tippen til dannelsen (anterograd transport)
og returnerer ”turnover” produkterne fra tippen og tilbage til basen (retrograd transport).
IFT systemet består af anterograd (kinesin-2) og retrograd (cytoplasmic dynein 2) motor
komplekser samt ca. 17 forskellige IFT partikel proteiner, delt i to store komplekser
kaldet A og B. De molekylære mekanismer hvorved disse forskellige komponenter bliver
koordineret og reguleret ved flagellets base og tip er usikkert.
Den encellede grønne alge Chlamydomonas reinhardtii er en veletableret
modelorganisme til studiet af cilier og IFT. IFT ”turnaround” i flageltippen involverer:
inaktivering/nedregulering af kinesin-2, aktivering/opregulering af cytoplasmisk dynein 2,
aflastning af flagellære byggesten og lastning af flagellære ”turnover” produkter. Det har
tidligere været vist at EB1, et lille MT plusende associeret protein (+TIP), lokaliserer til
flagellets tip i Chlamydomonas hvor det interagerer med IFT172 muligvis i reguleringen
af IFT partikel ”turnover”. Formålet med dette projekt var at karakterisere og identificere
flagel tip proteiner, der formodes at spille centrale roller i IFT regulering og/eller
ciliedannelse og funktion. Til dette brugte jeg Chlamydomonas som en modelorganisme
og benyttede to forskellige strategier. Først, ved at bruge et antistof genereret mod det
lille nukleotidbindende protein 1 (Nubp1) før starten af dette studie, viser jeg ved brug af
western blotting og immunofluorescens mikroskopi at Nubp1 er lokaliseret til flagellerne


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I Chlamydomonas og er specielt beriget ved flageltippen. Derudover ville jeg identificere
bindingspartnere til EB1 og IFT172 C-terminal ved brug af GST “pull-down” fra isolerede
flageller. Denne del af mit speciale arbejde blev gjort i samarbejde med Anna
Akhmanova og hendes gruppe i Rotterdam, Holland. Akhmanovas gruppe udførte GST
”pull-down”forsøgene i Chlamydomonas ved brug af isolerede flageller og GST-
EB1/IFT172 fusionsproteiner og identificerede formodede bindingspartnere ved
massespektrometri. Jeg klonede og karakteriserede derefter tre af de potentielle
EB1/IFT172 bindingspartnere der var blevet identificeret.


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Abbreviations
+TIP Plus-end tracking protein
AcTub Acetylated alpha-tubulin
Arf ADP-ribosylation factors
Arl Arf-like protein
BBS Bardet-Biedl syndrome
BCIP/NBT 5-bromo-4-chloro-3-indoylphosphate/Nitroblue tetrazolium
BSA Bovine serum albumin
C. reinhardtii Chlamydomonas reinhardtii
cDNA Complementary DNA
CGC Chlamydomonas Genetics Center
CrNubp1 Chlamydomonas reinhardtii Nubp1
DDT Dithiothreitol
DMSO Dimethylsulfoxide
EB End binding protein
E. coli Escherichia coli
EtOH Ethanol (CH3CH2OH)
FAP20 Flagella Associated Protein 20
GTP Guanosine triphosphate
GST Glutathione S-transferase
IC Intermediate chain
IFM Immunofluorescence Microscopy


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IFT Intraflagellar transport
Ig Immunoglobulin
KCBP Kinesin-like Calmodulin binding protein
KIF Kinesin superfamily protein
MAP MT-associated protein
MeOH Methanol (CH3OH)
MBP Maltose binding protein
MmNubp1 Mammalian Nubp1
mRNA Messenger RNA
MTOC MT organizing center
MT Microtubule
NIH3T3 National Institute of Health 3T3
Nubp1 Nucleotide-binding protein 1
OD Optical densities
PBS Phosphate buffered saline
PCD Primary cilia dyskinesia
PFA Paraformaldehyd
PKD Polycystic kidney disease
PCR Polymerase chain reaction
RPE Retinal pigment epithelial
RT-PCR Reverse transcription polymerase chain reaction
SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis


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SLB Selective Lhx3/4 Lim-homeodomain transcription factor
binding protein
SN Supernatant
SOFA Site of axonemal severing
TAE Tris-acetate-EDTA
TAP Tris-acetate-phosphate
TBS Tris buffered saline
TBST TBS Tween-20
TFIIB Transcription factor IIB
WB Western blot analysis


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1. Aim of study
1.1. Introductory remarks
Motile and the non-motile primary cilia are assembed and maintained by intraflagellar
transport (IFT), a highly conserved bidirectional microtubule-based transport system.
IFT transports flagellar precursors from the flagellar base to the tip for assembly
(anterograde transport) and returns turnover products from the tip back to the base
(retrograde transport). The IFT system consists of anterograde (kinesin-2) and
retrograde (cytoplasmic dynein 1) motor complexes, and ca. 17 different IFT particle
proteins separated in two large complexes, A and B. The molecular mechanisms by
which these different components are coordinated and regulated at the flagellar base
and tip are unclear. The molecular mechanisms of IFT turnaround in Chlamydomonas
involves: inactivation/down-regulation of kinesin-2, ativation/upregulation of cytoplasmic
dynein 2, unloading of flagellar precursors, and the loading of flagellar turnover
products. EB1 is a MT plus-end tracking protein (+TIP) and localizes to the flagellar tip
in Chlamydomonas reinhardtii (Pedersen et al., 2003), where it interacts with IFT172
possibly regulating IFT particle turnover (Pedersen et al., 2005).
1.2. Specific aims
The aim of this project was to use Chlamydomonas reinhardtii as a model, to identify
and characterize novel flagellar tip proteins, that is presumed to play central roles in the
building of the cilia as well as the regulation of cilia mediated signal transduction and the
cell cycle. First, based on the results for CrEB1 in the flagella proteome analysis
(Pazour et al., 2005), CrNubp1 was chosen, based on the fact that it had the same
properties as CrEB1 in having fex peptides and all in the membrane plus matrix
fractions. Furthermore, the mouse kinesin-14 family member KIFC5A is a minus-end-
directed kinesin involved in regulation of centrosome duplication and the cell cycle.
KIFC5A interacts directly with nucleotide-binding protein 1 (Nubp1) and the related
protein Nubp2, and inactivation of either KIFC5A or Nubp1 in mouse fibroblasts results
in the presence of supernumerary centrosomes and an increase in the proportion of bi-


12
and tri-nucleated cells (Christodoulou et al., 2006). Prior to the onset of this project, a
Chlamydomonas homolog of Nubp1 was cloned and an antibody generated against this
protein. My aim was to test this antibody and potentially characterize the protein.
Secondly, as an alternative approach to identifying novel tip proteins, I set out to
identify binding partners of C. reinhardtii EB1 and IFT172 C-terminus using GST pull-
down of isolated flagella from C. reinhardtii cells. This part of my thesis work was done
in collaboration with Anna Akhmanova and her lab in Rotterdam, The Netherlands.
Akhmanovas group executed the GST pull-down experiments in Chlamydomonas using
isolated flagella and GST-CrEB1 fusion protein and identified putative binding partners
by mass spectrometry. My aim was to clone three of the potential EB1/IFT172 binding
partners identified and retest this potential binding as well as to potentially a
characterized them.


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2. Introduction
2.1. Cilia and flagella structure
Cilia and flagella (the terms are equivalent1
) are long, thin organelles projecting like hair
from the surfaces of most eukaryotic cells, where they play important motile and
sensory functions (Christensen et al., 2007). The core of these organelles is composed
of a microtubule (MT)-based skeleton called the axoneme. The axoneme extends from
a modified centriole, the basal body, which anchors the axoneme in the cell. The
axoneme is surrounded by an extension of the plasma membrane continuos of the cell
body but the flagellum membrane is selectively different from the cell membrane in
overall composition, containing a different complement of membrane receptors and ion
channels (Christensen et al., 2007; Satir and Christensen, 2007). The axoneme is
composed primarily of MTs which are hollow cylinders built of heterodimers of α- and ß-
tubulin. The heterodimers bind together in a head-to-tail manner to form a protofilament
and the protofilaments bind side-by-side to form the MT wall. In this way the MTs
become polarized, with ß-tubulin at the fastest growing end called the plus end, and the
slowest growing end, the minus end, finishing with α-tubulin (reviewed in Desai and
Mitchison, 1997). MTs are very dynamic and are constantly polymerizing and de-
polymerizing. This occurs preferentially at the plus end, while the minus end is less
dynamic. The MT plus-ends are highly unstable switching rapidly between growth,
pause and shrinkage. This phenomenon is known as dynamic unstability and can be
modulated by MT-associated proteins (Howard and Hyman, 2003). The axoneme
consists of MT doublets, an A and B tubule, where the A tubule is a complete cylinder of
13 protofilaments and the B tubule an incomplete cylinder consisting of 10
protofilaments, attached to the A tubule (Figure 2.1.E). In the axoneme the MTs are
arranged such that the minus ends are embedded in the basal body while the plus ends
are oriented towards the tip of the cilium (Allen and Borisy, 1974). This means that the
axoneme is assembled at and constantly turning over at its tip which requires continous
transport of axonemal precursors from the cell body to the tip (Marshall and

1 The terms will be used interchangeably throughout this thesis.


14
Rosenbaum, 2001; Rosenbaum and Witman, 2002). In general however, axonemal MTs
are very stable: stable MTs are characterized by acetylation and detyronisation of α-
tubulin, which is important for cilia maintenance and function. Further, antibodies
against acetylated or detyrosinated tubulin are frequently used as markers for cilia and
flagella (Poole et al., 2001).
Cilia and flagella are classified, depending upon their axonemal structure, as
motile cilia or non-motile cilia. Motile cilia and flagella contain an axoneme with nine
outer doublet MTs, held together by nexin links, as well as a central pair of MT. They
therefore have a ”9+2” structure (Figure 2.1.E, left). The motile cilia usually play a role in
moving fluids over a cell layer or in movement of single cells (Marshall and Kintner
2008; Ginger et al., 2008), and contain accesory components involved in motility,
including outer and inner dynein arms and radial spokes (Figure 2.1.E, left). Motile cilia
(”9+2”) can be found in multiple copies per cell like in the respiratory epithelia (Figure
2.1, A), mammalian oviduct and brain ventricles. Here they are designed to move the
fluid and mucous overlaying the ciliated epithelium by the coordinated beating of the
cilia. When motile cilia (”9+2”) are found in one or two copies per cell e.g in mammalian
sperm cells (Figure 2.1.B) and in the green alga Chlamydomonas reinhardtii (Figure
2.1.C; see section 2.8), they are often known as flagella. Here the flagella are important
for the movement of the cell (Marshall and Kintner 2008; Ginger et al., 2008).
Non-motile cilia, also known as primary cilia, only exist in one copy per cell and
are present in vertebrate cells when these are in growth arrest (Schneider et al., 2005;
Figure 2.1.D). Their axoneme structure consists of a ”9+0” structure, which means that
they lack the central pair of MT. Furthermore they also lack the accessory components
involved in motility, that is seen in the motile cilia (Figure 2.1.E). Primary cilia have been
shown to be involved in coordination and regulation of a variety of crucial cellular and
developmental processes (Christensen et al., 2007). Modified primary cilia also exist
and are present on differentiated cells of the eye and olfactory organs and are essential
for the senses of sight, equilibrium and hearing (Singla and Reiter, 2006).
However there are examples of cilia, which do not clearly fit into either of these
two groups. The nodal cells of developing mammalian embryos have cilia sharing


15
features from both motile as well as primary cilia. These cilia have a ”9+0” structure, but
also posses outer arm dyneins. They generate a propeller-like motion that creates a
directional flow across the node required for establishment of the left-right asymmetry
axis (Hirokawa et al., 2006). And a novel 9+4 axoneme with four central MTs, have also
been identified on the notochordal plate of the rabbit embryo, thus indicating that some
degree of variation of axonemal structures exist (Feistel and Blum, 2006).
Figure 2.1: Cilia and flagella. A: Motile cilia on lung epithelia
(http://www.newscientist.com/data/images/ns/cms/dn11602/dn11602-2_585.jpg). B:
Spermatozoa approaching an egg (http://z.about.com/d/civilliberty/1/5/u/-/-/-/spermegg.jpg).
C: Scanning electron micrograph of the unicellular, biflagellated green alga,
Chlamydomonas reinhardtii (Pan et al., 2005). D: Scanning electron micrograph of renal
epithelial cells in a kidney collecting tubule. Each cell has a primary cilium (Ci) (Pan et
al., 2005). E: Axonemal structure. Cross section of motile 9+2 cilia (left) and immotile
9+0 cilia (middle). The axoneme is constructed of 9 doublet MTs connected via nexin
links. Motile cilia also have a central placed MT-pair. The basal body consists of triplet
MTs and no central pair (right). Axoneme figure modified from Dawe et al., 2007.


16
The ciliary axoneme is anchored to the basal body, which is a modified centriole. The
basal body differs in structure from the ciliary axoneme, by consisting of MT triplets
(Figure 2.1.E, right). Separating the membrane compartments of the cilia and the cell
body, at the ciliary base, is a region known as the ”ciliary necklace” or ”ciliary pore”,
which is connected by fibers to the transition zone of the basal body (Figure 2.2; Gilula
and Satir, 1972, see section 2.3).
2.2. Ciliopathies
Cilia are on almost every cell in the human body, where they play important motile and
sensory functions (Christensen et al., 2007), and it is therefore not surprising that
various human disorders can be related to defects in cilia. The phenotypes of these
disorders, the so-called ciliopathies, reflect the many roles cilia play in the human body.
Some examples are listed in table 2.1 (Badano et al., 2006; Marshall, 2008; Pan, 2008).
Defects are seen in both motile cilia and the non-motile primary cilia and the diseases
can either be linked to completely missing cilia or defects in or mis-localization of ciliary
proteins.
Defects in ciliary motility can lead to immotile cilia syndrome, also known as
primary cilia dyskinesia (PCD) (Bisgrove and Yost, 2006). The disease can be caused
by defects in multiple proteins involved in motility such as the dynein arms, the radial
spokes or the central pair MTs, and thereby only affects motile cilia (Afzelius, 2004). It
was a study of Chlamydomonas motility mutants defective in dynein that facilitated the
first identification of the genetic basis for PCD in patients (Pennarun et al., 1999).
Defects in motile cilia can also cause hydrocephalus (accumulation of water in the
brain) and altered left-right axis patterning during embryonic development as well as
infertility in male patients (Afzelius, 2004). Defects in the non-motile primary cilia can
lead to diseases caused by defects in signalling or assembly of the cilium. Examples
are polycystic kidney disease (PKD), Bardet-Biedl syndrome (BBS), polydactyly, obesity
and other more rare diseases. It was the study of flagellar assembly in Chlamydomonas
mutant ift88 that provided the first link between PKD and cilia (Pazour et al., 2000).


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Human disease Gene Cellular function Protein
localization
Disease pathology
PCD DNAH5
DNAI1
Ciliary motility Outer dynein
arms
Respiratory infections,
anosmia, male
infertility, otitis media
and situs invertus
Meckel-Gruber
syndrome
Cep290
MKS1;
MKS3
For Cep290
unknown;
Ciliogenesis
Basal body
and IFT
complexes
Brain malformation,
polydactyly, kidney
and liver cysts
PKD PKD1-2;
PKHD1
Mechanosensing;
PKHD1 unknown
Cilia and
basal body
Polycystic kidney
Nephronophtisis NPHP!-5 Uncertain Basal body
and cilia
Kidney cysts, liver
fibrosis, retinal
dysplasia
Joubert
syndrome
Central nerve system
abnormalities, kidney
cysts, brain and retina
malformations
Retinitis
pigmentosa
RPGR Retinal transport Basal body Retinal degeneration
BBS including BBS1-12 Ciliogenesis Basal body
and IFT
complexes
kidney cysts, obesity,
anosmia, retinal
dystrophy, male
infertility, situs
invertus, diabetes
Oral-facial-
digital syndrome
type I
OFD1 Ciliogenesis Basal body Malformations of the
face, oral cavity and
digits, kidney cysts
Table 2.1. Human ciliary disease genes and their cell biological functions. Table
modified from Marshall, 2008; D’Angelo and Franco, 2009.
2.3. Ciliogenesis and the cell cycle
As mentioned above the ciliary axoneme is anchored to the basal body, which is a
modified centriole, and assembly and disassembly of the cilia is therefore tightly
coupled to centriole duplication and the cyclic nature of the centrioles during the cell
cycle. This means that the formation of a primary cilium, ciliogenesis, is a regulated
process and closely connected to the cell cycle in proliferating cells forming a primary
cilium (Figure 2.2; Quarmby and Parker, 2005). The cilium is assembled during G1 by a


18
process called intraflagellar transport (IFT), after docking of the centrosome at the
plasma membrane and formation of the ciliary necklace (see section 2.3). The cilium is
most abundant in G0, and retracted in many cells at the entry into mitosis. Throughout
the cell cycle, the centrosome functions as a MT organizing center (MTOC), from where
the spindle poles are formed during mitosis and the primary cilium is nucleated during
growth arrest (G0) (Doxsey, 2001; Santos and Reiter, 2008). Centrosomes only
duplicate once per cell cycle and failure to do so correctly can result in e.g. multipolar
mitotic spindles and chromosomal missegregation. Several centrosomal proteins have
been determined to be essential for assembly of vertebrate primary cilia (Pedersen et
al., 2008). Others are linked to both cell cycle progression and resorption of the cilium
(Santos and Reiter, 2008). Of note is the mitotic regulatory kinase aurora A which
interacts with an adhesion scaffolding protein to control cilia disassembly (Santos and
Reiter, 2008). Aurora A is a member of Ipl family of kinases, and is modestly related to
CALK, a kinase involved in Chlamydomonas flagellar retraction (Pan et al., 2004) and
overactivity of aurora A and HEF1 has been associated with supernumerary
centrosomes and multipolar spindles (Pugacheva and Golemis, 2005).
2.4. Intraflagellar transport
The structure of cilia and flagella presents a transport problem since there is no protein
synthesis in the ciliary compartment. Cilia and flagella are assembled at the distal tip
(Johnson and Rosenbaum, 1992; Marshall and Rosenbaum, 2001), and therefore the
building blocks, which are synthesized in the cell body, must be transported to the tip to
assemble and maintain the flagella. This is done via IFT, a process essential for
assembly and maintenance of cilia and flagella (Kozminski et. al., 1993; Rosenbaum
and Witman, 2002; Cole, 2003). IFT was first observed in Chlamydomonas reinhardtii,
by Joel Rosenbaum´s group in 1993 as a transport system unrelated to ciliary beating
(Kozminski et. al., 1993), and it was later shown that IFT is an evolutionary conserved
process for building and maintaining cilia and flagella in such evolutionary distant
organisms as Caernorhabditis elegans and humans (Cole et al., 1998; Rosenbaum and
Witman, 2002).


19
Figure 2.2. Assembly and disassembly of primary cilia in the vertebrate cell cycle.
In most cells, primary cilium formation first occurs during G1, as the mother centriole
docks to the membrane. Assembly is mediated by IFT-dependent addition of ciliary
precursors as the cilium extends directly from the mother centriole’s triplet MTs. During
G1 and G0, the cilium functions as a cellular antenna. In the S-phase, the centrioles and
DNA replicate, and at entry to G2, the cilium is disassembled, so that the matured
centrioles can be ready for mitotic spindle formation. Once mitotic (M) cell division is
complete, the centrioles can proceed to ciliary re-assembly in G1. Figure from Pedersen
and Rosenbaum, 2008.


20
IFT is a bidirectional process that involves the movement of large protein complexes,
known as IFT particles, by two MT motor complexes responsible for anterograde (base
to tip) and retrograde (tip to base) transport. Cargo, such as building blocks and
turnover products, is coupled to the IFT particles, which are either classified as complex
A or complex B. Proteins destined for the cilium are assembled with the IFT particles
and motor complexes near the transition fibers, at a docking zone (Figure 2.3; Gilula
and Satir, 1972; Deane et al., 2001) and in this way enter the cilium. The transition
zone, also known as the “ciliary pore”, has been hypothesized to function as a barrier
controlling access of molecules to the cilium. The ciliary pore is thought to function, not
unlike the nuclear pore, as a regulated gate of entry where ciliary precursors and IFT
proteins accumulate prior to entering the ciliary compartment (Rosenbaum and Witman,
2002). Proteins destined for the cilium have signal targeting motifs, locating them there,
e.g. has the N-terminal RVxP motif been implicated in the localization of polycystin-2 to
the ciliary membrane (Geng et al., 2006). After assembly, the anterograde motor moves
along the B-tubules just underneath the ciliary membrane. When the complex reaches
the ciliary tip the cargo is unloaded and the IFT complex is reorganized. The retrograde
motor complex then transports new cargo back to the cytoplasm (Figure 2.3). Only one
motor complex is activate at a time and therefore the other is transported as cargo in an
inactivated form (Pedersen and Rosenbaum, 2008).
2.4.1. Anterograde IFT
Anterograde transport is reliant on kinesin-2 motor proteins, which can exist as either a
heterotrimeric kinesin-II or a homodimeric kinesin-2 (see section 2.7) In
Chlamydomonas, the heterotrimeric kinesin-II consists of two motor subunits FLA10 and
FLA8 of 90 and 85 kDa. respectively, and the non-motor subunit FLA3 (100 kDa),
known as the kinesin-2-associated protein or KAP (Kozminski et al., 1995; Scholey,
2003; Miller et al., 2005; Mueller et al., 2005). In human and mouse the motor domains
are called KIF3A and KIF3B, respectively. Furthermore a third motor subunit called
KIF3C exists and has been found to associate with KIF3A (Scholey, 2008; Pedersen et
al., 2008). Kinesin-II associates with IFT particles at the transition zone and this huge


21
motor complex transports axonemal precursors and other cargo molecules to the ciliary
tip.
Anterograde IFT has been extensively studied in Chlamydomonas. Analysis of a
temperature sensitive Chlamydomonas mutant, fla10ts
, which has a temperature-
sensitive mutation in the FLA10 gene (Adams et al., 1982; Vashishta et al., 1996), gave
the first indication that kinesin-II is required for anterograde IFT. When these
Figure 2.3: Assembly of cilia via intraflagellar transport (IFT). Ciliary proteins are
transported in Golgi-derived vesicles along cytoplasmic MTs to the ciliary base. Here
the ciliary proteins enter the cilium via the “ciliary pore” and the proteins are transported
anterogradely along the axoneme by kinesin-II. Ciliary turnover products are, in turn,
transported retrogradely along the ciliary axoneme by cytoplasmic dynein 2 for recycling
or degradation in the cytoplasm. From Pedersen and Rosenbaum, 2008.


22
mutant cells were placed at the restrictive temperature of 32°C, IFT ceased and the
flagella began to shorten. If the flagella were isolated from the mutant at the restrictive
temperature, new flagella failed to form. These experiments have shown that kinesin-II
and IFT are required for assembly and maintenance of the flagella. In Chlamydomonas,
kinesin-II is the only anterograde motor, but this is not the case for C. elegans, where a
homodimeric kinesin-2 consisting of OSM3, has been observed to play a role in
anterograde IFT, in part by working in concert with kinesin-II (Scholey et al., 2004).
Kinesin-II is the core anterograde IFT motor in virtually all ciliary systems studied to
date. However other accesory motors may cooperate with it, as seen with OSM3
(Scholey, 2008).
2.4.2. Retrograde IFT
Retrograde IFT is motored by an isoform of cytoplasmic dynein called cytoplasmic
dynein 2, previously known as cytoplasmic dynein 1b (Figure 2.4; Pfister et al., 2005;
Pedersen and Rosenbaum, 2008). Dyneins are minus-end directed multiprotein
motorcomplexes consisting of one or more heavy chains and several associated
proteins (Pedersen et al., 2008). In Chlamydomonas the motor complex consists of a
heavy chain, DHC1b (belonging to the AAA+ family of ATPases; Pazour et al., 1999;
Asai and Koonce, 2001), a light intermediate chain, D1bLIC (Hou et al., 2004), an
intermediate chain (IC)/WD repeat protein, FAP133, which may be specific for motile
cilia (Rompolas et al., 2007) and a light chain, LC8/FLA14 (Figure 2.4; Pazour et al.,
1998). In human and mouse, a motor heavy chain, DHC2, a light intermediate chain
D2LIC, and an intermediate chain, WD34, have been identified. However, the precise
function of the individual subunits of the cytoplasmic dynein 2 complex during
retrograde IFT is still unclear (Pedersen and Rosenbaum, 2008).
The cytoplasmic dynein 2 motor subunit, DHC2, was originally identified as a
dynein heavy chain in sea urchin embryos (Gibbons et al., 1994), and has subsequently
been studied further in Chlamydomonas and C. elegans (Pazour et al., 1999; Signor et
al., 1999).


23
Figure 2.4: Chlamydomonas cytoplasmic dynein 2.
The heavy chains bind the outer double MT, and the
light chains bind cargo. Modified from Rompolas et al.,
2007.
2.4.3. Tip turnaround
Both anterograde and retrograde IFT seem to occur at a constant rate along the cilium
(Kozminski et al., 1993), with sligth pauses at the base and tip, so the main points of
regulation of IFT are presumably at the ciliary base and tip (Pedersen and Rosenbaum,
2008). Cilia and flagella are assembled and continously turnover at their distal tip
(Johnson and Rosenbaum, 1992; Marshall and Rosenbaum, 2001). IFT transports
flagellar precursors from the flagellar base to the tip for assembly and returns turnover
products from the tip back to the base. The molecular mechanisms by which these
different components are coordinated and regulated at the flagellar base and tip are
unclear. The molecular mechanisms of IFT turnaround in Chlamydomonas involves: the
inactivation/down-regulation of kinesin-2, activation/upregulation of cytoplasmic dynein
2, unloading of flagellar precursors, and the loading of flagellar turnover products
(Pedersen and Rosenbaum, 2008). The timing and mechanisms of these events are
unknown, although some clues have recently emerged.
Regulation of kinesin-2 motor activity appears quite complex involving a variety
of different regulatory mechanisms (e.g. phosphorylation, comformational changes,
tubulin modifications) and molecules. To mention a few, analysis of mutants or
biochemical inhibitors affecting ciliary length, have revealed a number of kinases as


24
potential regulators of kinesin-II activity, including MAP kinases and NIMA-related
kinases (Pedersen and Rosenbaum, 2008). Also, kinesin-II activity may be regulated via
conformational changes in the KAP subunit, because KAP is required for localization of
kinesin-II at the flagellar base as well as for movement of the motor complex along
flagella in Chlamydomonas (Mueller et al., 2005). The mechanisms by which
cytoplasmic dynein 2 is regulated are virtually unknown. This could be due to the fact
that it has been difficult to purify biochemically, and the complex may contain additional
unidentified subunits (Pedersen and Rosenbaum, 2008).
At the flagellar tip, IFT particle turnover seems to be regulated by IFT172
(Pedersen et al., 2005; Tsao and Gorovsky, 2008) possibly in conjunction with the small
MT-associated protein EB1 (Pedersen et al., 2005), which localizes to the flagellar tip
and basal bodies in Chlamydomonas (Pedersen et al., 2003; also see section 2.4.4.1
and 2.5).
2.4.4 IFT particle polypeptides
Associated with to kinesin-II and cytoplasmic dynein 2 are IFT particles, which have
multiple protein-protein interaction motifs serving as docking sites for cargo proteins,
such as ciliary building blocks (Cole, 2003; Blacque et al., 2008;). IFT particle proteins
were first identified in, and isolated from Chlamydomonas flagella, using the fla10ts
mutant (Kozminski et al., 1993; Piperno and Mead, 1997; Cole et al., 1998; Also see
section 2.2.1). Sucrose density gradient centrifugation was used to fractionate the
membrane plus matrix, allowing comparison of flagellar proteins, extracted under mild
conditions (Piperno and Mead, 1997; Cole et al., 1998). This led to the identification of
approximately 17 different IFT particle proteins which can bee separated into two large
complexes, A and B (Cole et al., 1998; Piperno et al., 1998). Cloning and sequencing of
Chlamydomonas IFT particle polypeptide genes showed that both complex A and B
components have several domains and amino acid repeats typically involved in
transient protein-protein interactions (Cole, 2003). The IFT particle polypeptides all have
apparent molecular masses between 20 and 172, and are named IFT20 through IFT172
(Cole et al., 1998). Complex A comprises the following IFT particle proteins: IFT144,


25
IFT140, IFT139, IFT122A, IFT122B, and IFT43 and their overall function is primarily
associated with retrograde IFT. Complex B comprises the following IFT particle
proteins: IFT172, IFT88, IFT81, IFT80, IFT74/72, IFT57 (also known as IFT57/55),
IFT52, IFT46, IFT27, IFT25, IFT22 and IFT20. Complex B is required for anterograde
IFT and loss of any of the complex B proteins results in shortened or absent cilia. For
some complex B proteins, additional or specific functions related to flagellar assembly
have been described. For example, IFT20 is involved in transport of vesicles from the
Golgi to the ciliary base (Follit et al., 2006; Omori et al., 2008), IFT27 functions as G-
protein in the cell cycle (Qin et al., 2007), IFT46 is involved in transport of outer dynein
arms into the flagella (Hou et al., 2007), and IFT172 functions as a regulator of the
transition from anterograde to retrograde IFT in the tip of the cilia (Pedersen et al.,
2005; Tsao and Gorovsky, 2008).
2.4.4.1. IFT172
IFT172 is encoded by FLA11 and is the protein of complex B with the highest molecular
mass (172 kDa) (see section 2.4.4; Cole et al., 1998). The Chlamydomonas fla11
mutant has a point mutation in IFT172, which results in short or missing cilia as well as
accumulation of IFT particles in the ciliary tip (Pedersen et al., 2005). IFT172 has been
shown to contain a N-terminal WD repeat domain (WDD) composed mainly of β-sheets
and a C-terminal repeat domain (RPD) composed mainly of α-helices. In between the
RPDs a LIM interaction domain (LIM-ID) is located (Figure 2.5; Pedersen et al., 2005;
Tsao and Gorovsky, 2008). These structures have been shown to be very conserved
among different organisms and are involved in protein-protein interaction. Studies of
IFT172 in Tetrahymena, where the different domains had been selectively deleted,
showed that both the N- and C-terminal domains are essential for localization of IFT172
to cilia and for the assembly of cilia. A mutant with a partially truncated C-terminal
accumulated IFT particles in the ciliary tip, indicating failure of motor switching or
retrograde transport (Tsao and Gorovsky, 2008).
The IFT172 orthologue in rats is called SLB (Selective Lhx3/4 Lim-homeodomain
transcription factor Binding protein), and the LIM-binding domain has been shown to


26
interact speciﬁcally with members of the LIM homeodomain family of transcription
factors Lhx3 and Lhx4. Binding inhibits Lhx3 and Lhx4, indicating a possible role of
IFT172 as a transcription regulator (Howard and Maurer, 2000).
Mutant experiments in C. elegans have shown that mutations in the IFT172
orthologue OSM-1 gives a defect in sensory cilia (Perkins et al., 1986; Bell et al., 2006).
Furthermore, a screening in zebrafish mutants with kidney cysts, identified a mutation in
IFT172 (Sun et al., 2004). Finally, IFT172 dissociates easily from the rest of the IFT
complex B, which could indicate that it is in the periphery of the complex and thereby
ideally could be positioned to play a regulatory role in IFT (Pedersen et al, 2005).

Figure 2.5: IFT172 in Tetrahymena. The beta-sheats and alpha-helices are known for
protein-protein interaction. LIM indicates, the Tetrahymena IFT172 domain homologous
to the LIM-transcription factor domain in rats (SLB/IFT172). Modified From Tsao and
Gorovsky, 2008.
2.5. EB1
EB1 is a MT plus-end tracking protein (+TIP) and belongs to one of the most conserved
families among the +TIPs, the EB family. EB proteins contain highly conserved N- and
C-terminal domains, which are separated by a less conserved linker sequence. The N-
terminal domain is necessary for MT binding and the C-terminal domain has a coiled-
coil region that mediates the parallel dimerization of EB protein monomers and at the
same time forms a surface for binding of various partners (Lansbergen and Akhmanova,
2006). EB1 is a relatively small MT-binding protein and it preferentially localizes to the
plus end of cytoplasmic MTs where it is involved in regulating MT dynamics.
Furthermore it is indirectly involved in linking the plus end with the cell cortex, mitotic


27
kinetochores and different cellular organelles by directing other MT-associated proteins
(MAPs) towards the plus tip. EB1 also localizes to the centrosomes and is required for
centrosomal MT anchoring. In addition, EB1 has been shown to localize to the ciliary tip
and the proximal part of the basal bodies in Chlamydomonas (Pedersen et al., 2003),
and centrosomal EB1 is required for assembly of primary cilia in mouse fibroblasts, by
Interacting with p150Glued
in cilia formation (Schrøder et al., 2007). However, the exact
mechanism by which this occurs is unknown. Vertebrates contain two additional EB1-
like proteins (EB2 and EB3), and it is possible that EB2 and/or EB3 also contribute to
ciliogenesis.
2.6. Nucleotide-binding protein 1 (Nubp1)
The Chlamydomonas protein Nubp1 (Figure 2.6), which was identified as part of the
Chlamydomonas genome sequencing project (Merchant et al., 2007), is homologous to
NBP1 in humans that belongs to the NUBP/MRP subfamily (Figure 2.7; Nakashima et
al., 1999). Nubp1 proteins are very conserved in different eukaryotes and contain a
MRP domain, a P-loop containing nucleotide triphosphate hydrolases (ATP/GTP-
binding) site, as well as an α- and ß-motif (see figure 2.6; 2,7; Nakashima et al., 1999).
In mammals, Nubp1 is closely related to Nubp2; however Nubp1 has a unique N-
terminal extension containing four cysteine residues, lacking in the shorter form, Nubp2
(Nakashima et al., 1999).
Figure. 2.6: Schematic presentation of Chlamydomonas Nubp1. It has highly
conserved domains belonging to the NUBP/MRP subfamily: The ATP/GTP binding
domain (Blue) and NUBP/MRP consensus pattern (Green) as well as the α-(red) and ß-
(orange) motif; Also see Figure 2.7 for conservation of the motifs in different eukaryotes.


28
Nubp1 proteins have extensive similarity to the prokaryotic division-site-
determining membrane ATPase protein MinD (Nakashima et al., 1999). In bacteria,
FtsZ, the bacterial homologue of tubulin, assembles to a cytoskeletal element, the Z-
ring, that recruit other proteins to carry out cytokinesis. The positioning of the Z-ring is
determined by a gradient of negative regulators. MinD together with the min operon
proteins MinC and MinE cooperatively position the Z-ring, thereby determining the
separation site for cell division (reviewed by Lutkenhaus 2007).
The Chlamydomonas Nubp1 homolog has not previously been characterized
biochemically or functionally and studies on Nubp1 homologs in other organisms are
also scarce. However, in mouse fibroblasts, Nubp1 has been shown to interact directly
with the related protein Nubp2 as well as the minus-end directed kinesin KIFC5A.
Inactivation of either KIFC5A or Nubp1 in mouse fibroblasts results in the presence of
supernumerary centrosomes and an increase in the proportion of bi- and tri-nucleated
cells (Christodoulou et al., 2006). Whether these phenotypes in any way are coupled to
cilia is unknown.
2.7. Kinesins
Kinesins constitute a superfamily of ATPase motor proteins that travel along MT tracks.
The family mediates diverse functions in the cell, including the transport of vesicles,
organelles, chromosomes and protein complexes (Hirokawa et al., 1998; Dagenbach
and Endow, 2004; Miki et al., 2005). The motor domain of the kinesin superfamily is
very conserved, and differences in the ca. 350 amino acid sequence is basis for the
classification of the motor proteins (Kashina et al., 1997). So far 17 different kinesin
families have been described (Wickstead and Gull, 2006) and a general kinesin
nomenclature was introduced in 2004 (Lawrence et al., 2004).
Usually a kinesin motor protein also comprises a regulatory neck domain
following the motor, and a tail region that interacts with cargo or other subunits (Figure
2.8). In contrast to the motor region, the tail region can be highly diverse among
kinesins, even within a family, and appears to bind cargo via adaptor or scaffolding
molecules (Hirokawa and Noda, 2008; Wickstead and Gull, 2006).


29
Figure 2.7: Multiple sequence alignment of Nubp1 homologs. The highly conserved
domains of the NUBP/MRP subfamily include: The ATP/GTP binding domain (blue) and
NUBP/MRP consensus pattern (green) as well as the α-(red) and ß-(orange) motif; See
Figure 2.6 for a schematic presentation of Chlamydomonas Nubp1. Data obtained from
Nakashima et al., 1999.


30
In most kinesins the motor domain is located in the N-terminus and such kinesins move
in the MT plus end direction whereas kinesins with the motor domain in the C-terminus
are minus end directed (Hirokawa and Noda, 2008). Normally kinesins are associated
as di- or trimers with the tails coiled together (Miki et al., 2001) and they walk in a cyclic
”hand-over-hand” manner of which many models exist. Basically, when the front motor
subunit binds ATP a conformational change displaces the weakly MT-interacting rear
head towards the MT-plus (or minus) end ahead of the other subunit and rebinds tightly
to ATP and the MT. The now rear head hydrolyses ATP causing a conformational
change releasing it from MT (Gennerich and Vale, 2009). I will shortly mention a few of
the kinesin families of interest to this thesis.
Figure 2.8. Schematic structure of conventional kinesin. Kinesin consists of a motor
domain (head), a coiled-coil stalk region, and a cargo binding region (tail). Figure from
Kikkawa, 2008.
2.7.1. Kinesin-2 family proteins
Kinesin-2 family members are known to participate in organelle transport, IFT and
spermatogenesis (Miki et al., 2005). Kinesin-II, the motor for anterograde transport, has
been described in section 2.4.1. However, kinesin-II also has non-cilia related functions
such as vesicle transport in neuronal axons (Hirokawa and noda, 2008)
Members of this family have not yet been described in fungi or higher plants that lack
cilia/flagella and sperm.


31
The kinesin-5 family proteins, also known as the BimC family, where given the family
number 5 because of the name of one of its most well known members, the mammalian
kinesin Eg5. The kinesin-5 family is the most conserved, monophyletic family and
kinesin-5 members are characterized by a characteristic BimC box domain (Kashina et
al., 1997). Kinesin-5 family members are found in mammals, yeast and higher plants.
They usually form homotetramers and are known to be mitotic motors functioning in
formation of the bipolar spindle during cell divison (Valentine et al., 2006). It is
hypothesized that they act in concert with minus-end-directed dyneins and other plus-
end-directed kinesins as well as serve to control the position of centrosomes and thus
play global roles in establishing and maintaining bipolar mitotic spindle structure. In all
known cases kinesin-5 family proteins prove to be localized to spindle MT (Kashina et
al., 1997). Mutations in the yeast bimC gene appear to block the separation of
duplicated centrosomes or spindle pole bodies resulting in the formation of defective
”monastral” mitotic apparati at early stages of mitosis. Furthermore, immunodepletion of
Eg5 in Xenopus oocytes causes defects in spindle formation at early stages of mitosis
(Kashina et al., 1997; Kapoor et al., 2000). Eg5 has also been shown to be expressed in
rodent postmitotic neurons. Here Eg5 is believed to be involved in organizing MT in the
devolping neurons (Ferhat et al., 1998). A specific inhibitor, called Monastrol, is known
to act specifically on the motor domain of human Eg5 arresting cells in mitosis (Cochran
et al., 2005).
Kinesin-14 family members are minus-end directed motors that cross-link MTs and play
key roles during spindle assembly. This family acts to regulate spindle length during
mitosis by cross-linking and sliding between parallel microtubules (Christodoulou et al.,
2006; Cai et al., 2009). Structurally Kinesin-14s have a conserved kinesin-like motor
domain at the C-terminus, a central coiled-coil stalk, and an N-terminal globular domain.
The mouse kinesin-14 member KIFC5A is involved in regulation of centrosome
duplication. Overexpression causes formation of aberrant, non-separated MT asters


32
and mitotic arrest in a promethaphase-like state. It is believed that the C-terminal minus-
end directed kinesins can produce forces that oppose the bimC-driven forces in the
mitotic spindle (Kashina et al., 1997), and knockdown of KIFC5A has been shown to
partly relieve the effect of the Eg5 inhibitor monastrol, indicating involvement in the
balance of forces determining the bipolar spindle during mitosis (Christodoulou et al.,
2006). KIFC5A interacts directly with Nucleotide-binding protein 1 (See section 2.6), and
inactivation of KIFC5A in mouse fibroblasts result in the presence of supernumerary
centrosomes and an increase in the proportion of bi- and tri-nucleated cells
(Christodoulou et al., 2006).
Chlamydomonas KCBP is a unique kinesin of the kinesin-14 family in that has a
calmodulin-binding domain. Close homologs are found in plants and there is also less
wellconserved member in sea urchins (Miki et al., 2005). Cytoplasmic dyneins are
lacking in plants and this could explain the abundance of this family member here, as
both are MT minus end-directed motors. Recently a KCBP kinesin has been shown to
localize to flagella and near the base of the flagella in Chlamydomonas. Although direct
functional data are lacking, this kinesin has been suggested to play a role in flagellar
assembly/disassembly as well as cell division (Dymek et al., 2006).
2.8. Chlamydomonas as a model organism for ciliary functions
Chlamydomonas reinhardtii is a motile single-celled green alga about 10 µm in diameter
and with two similar flagella of approximately 12 µm in length, which it uses for motility.
Chlamydomonas has become a model of great importance in the world of biology.
Chlamydomonas possesses properties from both the animal and plant kingdom,
showing similarity to animal cells by having centrioles and ﬂagella, but at the same time
also a relation to the plants, because it contains a chloroplast. These and other
properties make it the perfect model to study fundamental processes such as motility,
photosynthesis, cell cycle defects, responses to external stimuli such as light, and cell-
cell recognition (reviewed by Harris, 2001).


33
Chlamydomonas belongs to a very small group of model organisms where it is possible
to combine biochemical, genetic, and biological approaches to investigate the basic
biology and functions of ciliary and
basal body proteins (Badano et al.,
2006; Pan, 2008). Flagella of
Chlamydomonas are typical of
eukaryotic cilia and flagella, in that
they are composed of MT arranged
in the "9 + 2" structure (Figure 2.9).
Since different organisms solve
similar problems in similar ways,
studies on how MT assembly is
regulated in Chlamydomonas may
reveal mechanisms that are shared
by most other ciliated organisms.
IFT can be visualized in vivo without
the aid of fluorescence tagged
proteins and a large number of IFT mutants are available (Figure 2.9). Not surprisingly,
the protein components of the IFT machinery were first isolated and identified in
Chlamydomonas (Kozminski et al., 1993; Piperno and Mead, 1997; Cole et al., 1998;
section 2.4) and its flagella have been used extensively in the studies of IFT and other
cilia-related processes. Many known mutants of Chlamydomonas exist and make useful
tools for studying a variety of biological processes, including flagellar motility,
photosynthesis or protein synthesis. Since Chlamydomonas species are normally
haploid, the effects of mutations are seen immediately without additional timeconsuming
backcrossing (Figure 2.10; Pan and Snell, 2000). For instance a study of
Chlamydomonas mutants defective in dynein facilitated the identification of the genetic
basis for the disease PDC in human (Pennarun et al., 1999), and another study of a
mutant with defects in flagellar assembly (mutant ift88) provided the first link between
PKD and cilia (Pazour et al., 2000).


34
Figure 2.10: The Chlamydomonas cell cycle. 1-7: sexual reproduction 8: asexual
reproduction. 1. Gametogenesis is induced, when the N-source is removed. 2. Adhesion
of gametes of opposite mating types. 3. The cell walls are released and the mating
structures activated. 4. Fusion of mt+ fertilization tubule with mt- mating structure. 5.
Complete cell fusion. 6. Zygote (2n) maturation. 7. When the environment is optimal the
zygote undergoes meiosis/germination and new vegetative cells (1n) are formed. 8.
Cells undergo mitosis by resorbing the flagella and divide inside the cell wall of the
mother cell wall (not shown). For C. reinhardtii 2 mating types, mt+ and mt-, exist. The
figure was kindly provided by Ph.D. student Jacob M. Schrøder.


35
Chlamydomonas has a relatively short reproduction time of approximately 12-18 hrs,
depending on the temperature, light and media. Exposure to sunlight in an appropriate
medium produces uniform cultures containing large numbers of motile cells. The
Chlamydomonas cell cycle can be synchronized with cycles of light and dark, which is a
major technical advantage. Synchronization of the cells gives a higher yield of
flagellated cells for experimental purposes and the assembly and resorption of flagella
can more easily be studied, because the flagella are resorbed during entry into the
mitotic cycle and then reassembled after completion of the cycle. When
Chlamydomonas is grown with a 12:12, 14:10 or 16:8 light:dark cycle it can be fixed in
the G1 phase during the entire light
phase (Figure 2.11; Harris, 2001).
Chlamydomonas flagella can
readily be amputated and regrown,
observed, and measured. They can
be induced to gradually shorten
their flagella lengths, resulting in
complete loss, making it easy to
study the kinetics of flagellar
assembly and disassembly.
Chlamydomonas is one of the few
organisms from which cilia can be
isolated and purified in large
quantities, and at different stages of flagellar growth and shortening. A wide range of
chemical and physical stimuli can induce flagellation. The immediate response to an
acid shock produces intracellular acidification that induces an influx of calcium and
starts a signalling cascade resulting in activation of the severing machinery. The nine
outer doublet MT are broken at the distal end of the flagellar transition zone, the site of
axonemal severing (SOFA) (Sander and Salisbury, 1989; Quarmby and Hartzell, 1994;
Quarmby, 1996). Shedding of the flagella with a pH shock thus gives a very simple
biochemical method for isolating (and purifying) flagella.


36
By now, the genome sequence (Merchant el al., 2007), flagellar proteome
(Pazour et al., 2005), and flagellar transcriptome (Stolc et al., 2005) of Chlamydomonas
are known, which makes it easy to obtain the bioinformation needed for further research
with Chlamydomonas.
Since all organisms are related by evolution, the knowledge acquired from
studies of Chlamydomonas allows researchers to learn more about regulation of gene
expression in more complex plants and animals. Usually, finding the gene responsible
for a particular mechanism in human tissue without studying simpler model organisms is
nearly impossible. In this thesis, I have used Chlamydomonas as a model organism for
identifying and characterizing new proteins that localize to the flagellar tip, and which
may potentially be involved in regulating IFT and/or flagellar assembly/disassembly.


37
3. Results and discussion
3.1. Introductory notes
Axonemal MTs are oriented with their plus end towards the flagellar tip, where constant
assembly and turnover takes place (Johnson and Rosenbaum, 1992; Marshall and
Rosenbaum, 2001; Marshall et al., 2005). It has previously been shown that EB1, as
one of few proteins, localizes to the flagellar tip in C. reinhardtii (Pedersen et al., 2003).
CrEB1 interacts with IFT172 at the ciliary tip, where they may regulate IFT particle
turnover (Pedersen et al., 2005). In the known proteomic analysis of the C. reinhardtii
flagellum (Pazour et al., 2005), EB1 was represented with very few peptides (2 unique
peptides) and all in the membrane plus matrix fraction.
Prior to the onset of my project, to identify other potential flagellar tip proteins,
proteins also found in the flagella proteome with approximately the same peptides and
exclusively in the membrane plus matrix fraction were identified by the Pedersen
laboratory, and a protein homologous to mouse Nubp1 (CrNubp1) was selected as an
interesting potential flagellar tip protein, because mouse Nubp1 is known to interact with
KIFC5A, a minus end-directed kinesin of the kinesin-14 family (Christodoulou et al.,
2006). The N-terminal cDNA coding region of CrNubp1 (GenBank accession,
gi:159485046; nt 1-619) was cloned and sequenced by the lab, and a polyclonal
antibody against the N-terminus of CrNubp1 was produced prior to the onset of this
project. I affinity purified the antibody, tested it and used it in my project to characterize
CrNubp1. These results are presented in sections 3.2-3.5. In addition, I obtained two
different antibodies specific for mouse Nubp1 (provided by Niovi Santama, University of
Cyprus), and I used these antibodies for immunofluorescence microscopy (IFM)
analysis of mouse NIH3T3 cells to determine whether Nubp1 localized to primary cilia or
basal bodies in these cells (section 3.6).
As an alternative approach to identifying novel tip proteins, I set out to identify
binding partners of CrEB1 and IFT172 C-terminus using GST pull-down of isolated
flagella from C. reinhardtii cells. This part of my thesis work was done in collaboration
with Anna Akhmanova and her lab in Rotterdam, The Netherlands. Akhmanovas group


38
executed the GST pull-down experiments in Chlamydomonas using isolated flagella and
GST-CrEB1 fusion protein (Pedersen et al., 2005), and identified putative binding
partners by mass spectrometry. The flagella were isolated and purified by me (see
section 5.2) and the resulting data was analysed by my supervisor Lotte Pedersen. I
subsequently cloned and characterized three of the potential EB1/IFT172 binding
partners identified (section 3.7-3.10).
3.2. Chlamydomonas reinhardtii Nubp1 is present in the flagella
To characterize C. reinhardtii Nubp1 (CrNubp1), an antibody against the N-terminal part
of this protein was affinity purified and tested using western blotting (see section 5.10)
to see if it was present in the flagella. CrNubp1 is predicted to encode a 40.6 kDa
protein (Table 3.1). Immunoblot analysis with the affinity purified CrNubp1 antibody
detected a single ~55 kDa band in isolated C. reinhardtii flagella (Figure 3.1.A). The
apparent molecular weight of this protein is higher than predicted, which could be due to
the acidic nature of CrNubp1 (predicted pI of 4.95; Table 3.1) resulting in slower
migration during SDS-PAGE.
To compare the relative abundance of CrNubp1 in flagella and cytoplasm, blots
of wild type (CC-124) de-flagellated cell bodies were compaired with flagella isolated
from an equal number of cells (Figure 3.1.B). CrNubp1 is enriched in the flagella, similar
to IFT139 (IFT complex A protein). The majority of the cellular pool of CrEB1 is in the
cell body consistent with published data (Figure 3.1.B; Pedersen et al., 2003).
Table 3.1: Theoretical values for CrNubp1. The MW and pI values were calculated
from the polypeptide sequences using the pI/MW tool at Expasy.ch. ”Peptides” refers to
the number of peptides by which this protein is represented in the C. reinhardtii flagellar
proteome (Pazour et al., 2005).
MW (kDa) GenBank ID pI Peptides Amino acids
CrNubp1 40.6 gi: 159485046 4.95 1 1215


39
Figure 3.1: Presence of CrNubp in C. reinhardtii flagella. Western blot analysis
with affinity-purified CrNubp1 antibody detects a single band in wild type (CC-124)
isolated flagella (A). In (B) protein samples prepared from de-flagellated wild type
(CC-124) cell bodies and flagella were analyzed by western blot with antibodies
against IFT139 (IFT complex A protein), CrNubp1 or CrEB1. The bottom panel shows
a Coomassie-blue-stained gel run in parallel. Note that CrNubp1 is enriched in the
flagella, similar to IFT139, while the majority of the cellular pool of CrEB1 is in the cell
body.
Cell equivalents4
25 -
35 -
45 -
55 -
70 -
95 -
130 -
170 -
Coomassie
CrEB1
CrNubp1
IFT139
cell body flagella
BA
70 -
55 -
45 -
35 -
25 -
kDa
95 -
130 -
170 -
803 2 1 8 4 2 1


40
3.3. CrNubp1 localizes to the soluble membrane plus matrix
compartment
To determine where in the flagella CrNubp1 is present, wild type (CC-124) flagella were
disrupted by freezing and thawing in a detergent buffer. ATP was added, and the lysate
sucked through a 27 gauge needle, followed by centrifugation to sediment axonemes
and detergent-insoluble membranes (see section 5.2). This resulted in the release of all
of the total flagellar CrNubp1 into the supernatant/soluble fraction (Figure 3.2). This is
consistent with the finding that the one unique peptide of CrNubp1 found in the
Chlamydomonas flagellar proteome analysis was in the detergent-soluble membrane
plus matrix fraction (Pazour et al., 2005). Most of CrEB1 was also released in this
fraction. Upon further extraction of the pellet with high salt buffer, a small fraction of
CrEB1 was also associated with the salt extract (Figure 3.2). Thus, most of the flagellar
CrEB1 is soluble as consistent with previously published data (Pedersen et al., 2003;
Pazour et al., 2005). In contrast, several IFT components (IFT139, Fla10, IFT172,
D1bLIC) as well as the kinesin KCBP and Lis1-like protein CrLis1 were also present in
extracted axonemes in addition to the membrane matrix and high salt extract fractions
(Figure 3.2).


41
Figure 3.2: In fractionated wild
type (CC-124) flagella, CrNubp1
localizes to the soluble
membrane plus matrix
compartment. Flagella protein
samples from wild type cells were
fractionated as described in
materials and methods and
immunoblotted with antibodies
against IFT139 (IFT complex A
protein), KCBP (Kinesin-like
Calmodulin-binding protein; Dymek
et al., 2006), FLA10 (Kinesin-II
motor domain), CALK (Aurora
protein kinase; Pan et al., 2004),
IFT72 (IFT complex B protein),
D1bLIC (Cytoplasmic dynein 2 light
intermediate chain; Hou et al.,
2004), CrNubp1 (this study), CrLis1
(Lissencephaly protein Lis1;
Pedersen et al., 2007), and CrEB1
(Pedersen et al., 2003). Bottom
panel shows a Coomassie-blue-
stained gel run in parallel. Note that
CrNubp1 is in the membrane plus
matrix fraction only. Most of CrEB1
is also present in this compartment
consistent with previously published
results (Pedersen et al., 2003).


42
3.4. The flagellar level of CrNubp1 is unaffected by mutations
affecting assembly of the main axonemal substructures: outer dynein
arms, inner dynein arms, radial spokes and the central apparatus.
To determine if CrNubp1 is affected by the lack of axonemal components necessary for
flagella motility, flagella from different mutant strains were isolated and immunoblotted
with antibodies against CrNubp1, IFT139 (IFT complex A protein) and CrEB1 (Figure
3.3). CrNubp1 is found in flagella of all Chlamydomonas mutants examined here,
including those with flagella that lack radial spokes (pf14) and the central pair (pf18), as
well as mutants that lack the inner (ida1/ida4) and outer (oda2) dynein arms (Figure
3.3). Therefore CrNubp1 most likely does not localize to any of these axonemal
structures. This is consistent with the flagellar fractionation data indicating that CrNubp1
is only found in the soluble membrane plus matrix fraction of wild type (CC-124) flagella
(Section 3.3; Figure 3.2; Pazour et al., 2005). The same is seen for CrEB1 also
consistent with Pazours proteome analysis. Only a small band is visible for IFT139 in
the ida4 mutant strain (Figure 3.3). The explanation for this observation is unclear and
cannot be the lack of flagella inner dynein arms, because no effect is seen in the ida1
mutant. It is possible that the ida4 mutant strain harbors some additional mutation that
affects IFT139, but further experiments are needed to clarify this.
Figure 3.3: The flagellar level of
CrNubp1 is unaffected by mutations
in genes affecting motility-related
axonemal structures. Wild type
flagella (wt; CC-124) and flagella
isolated from different Chlamy-
domonas mutant strains were analyzed
by western blotting using antibodies
against CrNubp1, IFT139 and CrEB1,
as indicated. ida1 and ida4: lack inner
dynein arms; oda2: lacks outer dynein
arms; pf14: lacks radial spokes; pf18:
lacks entire flagellar central apparatus.
For details about these strains, see
www.chlamy.org.


43
3.5. CrNubp1 localizes to the basal bodies and the tip of the flagella
It was found by western blotting that CrNubp1 is present in the flagella, specifically in
the soluble membrane plus matrix compartment (Figure 3.1.A and Figure 3.2). To study
the localization pattern of CrNubp1 in whole C. reinhardtii wild type cells, IFM using the
polyclonal antibody directed against the N-term of CrNubp1 and a monoclonal anti-
acetylated alpha tubulin antibody to stain the flagella and basal bodies, was performed.
Using a methanol (MeOH) fixation method (see Section 5.11.1) this analysis showed
that CrNubp1 localized to the basal bodies in Chlamydomonas wild type cells (Figure
3.4). This result, however, shoul be interpreted with some caution because in contrast to
isolated flagella, western blotting of de-flagellated cell bodies or whole cells using the
CrNubp1 antibody failed to detect a band of the appropriate size (Figure 3.1.B and data
not shown), and therefore we do not know the specificity of the CrNubp1 antibody in the
cell body. However, similar analysis in mouse NIH3T3 cells strongly suggest that Nubp1
localizes to the basal bodies (see Section 3.6). In addition to possible basal body
localization of CrNubp1, in some cases, weak fluorescence was detected at the flagellar
tip on the MeOH fixed cells (data not shown).
To explore this possible tip localization further, IFM using an alternative fixation
protocol was performed. Interestingly, when cells were fixed using a fixation buffer with
Glutaraldehyde/NP40 (Lechtreck et al. 2009; Section 5.11.1) flagellar tip localization of
CrNubp1 in Chlamydomonas wild type cells was clearly observed (Figure 3.5). This is
consistent with the hypothesis that CrNubp1 is a flagellar tip protein and is also
consistent with my results indicating that CrNubp1 co-fractionates, at least in part, with
the known flagellar tip protein CrEB1 (see Section 3.3). Since mouse Nubp1 is known to
interact directly with the minus-end-directed kinesin KIFC5A (Christodoulou et al.,
2006), it is tempting to speculate that CrNubp1 similarly interacts with a minus-end-
directed kinesin at the flagellar tip in order to regulate flagellar disassembly and/or
transport of flagellar turn over products from the tip towards the cell body. However,
attempts to identify interaction between CrNubp1 and the known flagellar minus-end
directed kinesin KCBP (Dymek et al., 2006) were unsuccessful. It is of interest, though,


44
that minus-end-directed kinesins of the kinesin-13 family were identified at the flagellar
tip in Leishmania (Blaineau et al., 2007) and Giardia (Dawson et al., 2007).
Figure 3.4: IFM, using a methanol fixation method, shows basal body
localization of CrNubp1 in C. reinhardtii wild type (CC-124) cells. Cells were
grown at 20°C and subjected to IFM using a polyclonal antibody specific for
CrNubp1 (red). To detect the flagella and basal bodies an antibody specific for
acetylated alpha tubulin (green) was used. The IFM indicates that CrNubp1 (red)
localizes to the basal bodies of the cells. Asterisks mark the basal bodies, shown
enlarged in the insets.


45
Figure 3.5: IFM, using a Glutaraldehyde/NP40 fixation method, showing flagellar
tip localization of CrNubp1 in C. reinhardtii wild type (CC-124) cells. Cells were
grown at 20°C and subjected to IFM using a polyclonal antibody specific for CrNubp1
(red). To detect the flagella and basal bodies an antibody specific for acetylated alpha
tubulin (green) was used. The two bottom panels show the tips of flagella in focus. The
bottom panel in the middle was a control were no primary CrNubp1 antibody was
added. The IFM shows that CrNubp1 (red) localizes to the tip of the flagella, and no tip
localization is seen when no CrNubp1 primary antibody is added. The strong
fluorescence of the cell body is due to autofluorescence of the cells and was also
observed when no primary antibodies were added (not shown). Scale bar 10 µm.
CrNubp1!
No primary antibody!
AcTub! Merge!


46
3.6. Mammalian Nubp1 localizes to the centrioles and the nucleus in
NIH3T3 fibroblast cells
To study the localization pattern of Nubp1 in NIH3T3 mouse fibroblast cells, IFM using
two different antibodies against mouse Nubp1, generously provided by Niovi Santama,
University of Cyprus (Christodoulou et al., 2006), was performed. The first antibody is
an affinity-purified anti-peptide antibody made in guinea pig against the C-terminus of
Nubp1 and the second antibody is an affinity-purified antibody made in rabbit against
bacterially expressed recombinant Nubp1 (both unpublished). In addition, cells were
stained with a monoclonal anti-acetylated alpha tubulin antibody to stain the flagella and
basal bodies. Using the antibody against the C-term of Nubp1, the IFM analysis of
serum starved NIH3T3 mouse fibroblast cells showed that Nubp1 appeared to
concentrate mainly in the nucleus (Figure 3.6; Bottom panel; Data not shown). This was
also seen in cells in interphase (Data not shown). However, when using the antibody
against recombinant Nubp1, localization to the basal bodies was also observed (Figure
3.6; Three top panels). These results supports my observations in the IFM on
Chlamydomonas using CrNubp1 antibody (Figure 3.4) despite the fact that I was unable
to detect CrNubp1 in cell bodies by western blot analysis. It is possible that so little
Nubp1 exists in the cell so that it is not detectable with our particular antibody. Taken
together, the results indicate that Nubp1 is found at the flagellar tip (Section 3.5 ,Figure
3.5) as well as in the basal bodies (Section 3.5, Figure 3.4; Figure 3.6). What is the
function of CrNubp1 at these sites? As mentioned above, CrNubp1 at the flagellar tip
could function as a regulator of flagellar disassembly or transport between the tip and
the basal bodies by affecting minus-end-directed kinesins. In mouse 3T3 fibroblast cells,
Nubp1 has been shown to interact directly with Nubp2 as well as KIFC5A. The mouse
kinesin-14 family member KIFC5A is a minus-end-directed kinesin involved in regulation
of centrosome duplication. Inactivation of either KIFC5A or Nubp1 in mouse fibroblasts
results in the presence of supernumerary centrosomes and an increase in the
proportion of bi- and tri-nucleated cells (Christodoulou et al., 2006). The results
presented here suggest that Nubp1 at the flagella tip and basal bodies might be
important for regulating centriole duplication and cell cycle progression via modulation


47
of the activity of specific minus-end-directed kinesins. It has been shown that a member
of the kinesin-14 family, called kinesin-like Calmodulin-binding protein (KCBP), does
exist in Chlamydomonas flagella and is localized near the base of the flagella in
interphase. Although direct functional data are lacking, this kinesin has been suggested
to play a role in flagellar assembly as well as cell division (Dymek et al., 2006). I have
so far been unable to detect association between KCBP and CrNubp1 using
immunoprecipitation and MBP pull-down assays (data not shown), but it might be worth
to re-examine possible interactions between these proteins using alternative
approaches. In addition, it would be worthwhile to examine the localization of Nubp1 at
different stages of the cell cycle and to look for additional binding partners.
3.7. Identification of possible binding partners to CrEB1 and IFT172 C-
term in Chlamydomonas wild type (CC-124) flagella
To identify novel tip proteins, binding partners of IFT172 C-term and CrEB1 were
identified using Glutathione S-transferase (GST) pull-down of isolated flagella from wild
type (CC-124) cells. IFT172 plays a central role in the regulation of the transition
between anterograde and retrograde IFT at the tip of the flagellum, and a point mutation
in the C-terminus of IFT172 leads to accumulation of IFT particles at the flagellar tip
(Pedersen et al., 2005). IFT172 also interacts, at least indirectly, with CrEB1 (Pedersen
et al., 2005), which in turn is necessary for the formation of cilia in mouse fibroblast cells
(Schrøder et al., 2007). Finding binding partners to IFT C-terminus and CrEB1 might
therefore provide new insight into the mechanisms involved in IFT turn-over at the tip of
the flagella, as well as the regulation of flagellar MT elongation or disassembly.
GST pull-down experiments in Chlamydomonas using isolated flagella were
executed by Anna Akhmanovas group in The Netherlands. They ran the pull down
products on a SDS-PAGE gel (Figure 3.7) and identified putative binding partners using
mass spectrometry. The data was analysed by Lotte Pedersen, and a number of
potential binding partners of IFT172 C-term and CrEB1 in Chlamydomonas flagella
were identified (Appendix M). Three potentially interesting binding partners were chosen


48
for further analysis: an ARF-like protein (ARFA1A), a flagellar-associated protein
(FAP20) with homology to transcription factor IIB, and an Eg5-like kinesin motor protein.
Figure 3.6: IFM showing centriolar and nuclear localization of Nubp1 in growth-
arrested NIH3T3 fibroblast cells. Cells were starved for 48 hours and subjected to
IFM using an antibody specific for acetylated alpha tubulin (red) to detect the flagella
and centrioles. In the three top panels an antibody specific for mouse Nubp1, made in
rabbit, was used (green). In the bottom panel an antibody specific for the C-terminus of
mouse Nubp1, made in guinea pig, was used (green). Nuclei were stained with DAPI
(blue). Inserts: enlarged, shifted images of the centrioles and cilia. The IFM shows that
Nubp1 (green) localizes to the centrioles as well as the nucleus of the cells. Scale bar
10 µm.
AcTub! MmNubp1! Merge!DAPI!
Rabbitanti-Nubp1
Guineapig
anti-Nubp1


49
Figure 3.7: Coomassie-
stained gel showing the
products obtained by pull-
down assay of wild type
(CC-124) flagella with GST,
GST-CrEB1 and GST-
IFT172 C-term, respectively.
Image provided by Anna
Akhmanova.
The small ARF (ADP-ribosylation factors)-related GTPase, ARFA1A, was pulled
down with both GST-CrEB1 and GST-IFT172 C-term. The ARF protein family
compromises structually and functionally conserved members of the Ras superfamily of
regulatory GTP (Guanosine triphosphate)-binding proteins with many proposed
functions in mammalian cells, including the regulation of several steps of membrane
transport (Figure 3.8). Recent results suggests that several ARL (ARF-like) proteins may
be involved in different aspects of MT-dependent functions as well as activation of
phospholipase D (Kahn et al., 2005). Some ARLs have been implicated in ciliopathies:
mutations in the cilia gene ARL13B lead to Joubert syndrome (Cantagrel et al., 2008)
and ARL6 has been identified as one of the genes underlying Bardet-Biedl syndrome
(BBS) (Fan et al., 2004). Chlamydomonas ARFA1A is most closely related to the human
ARF members ARF1 and ARF3-5, which have been implicated in effects on Golgi and


50
endosome morphology (Kahn et al., 2005). In a parallel study by the Akhmanova and
Pedersen groups, ARF4 was identified as a possible binding partner of human EB1, and
other (unpublished) lines of evidence by our group suggest that transport of vesicles to
the ciliary compartment is impaired when EB1 is inactivated. The potential binding of
ARFA1A to EB1 or IFT172 is therefore very interesting.
Figure 3.8: Conserved domains of Chlamydomonas ARFA1A. ARFA1A belongs to
the family of small GTPases. The ARF family is a part of the superfamily Ras
GTPases. The ARFA1A sequence was blasted for conserved domains on
ncbi.nlm.nih.gov.
Flagella associated protein (FAP20) was also pulled down with both GST-CrEB1
and GST-IFT172 C-term. It belongs to the DUF667 superfamily highly similar to
vertebrate transcription factor IIB (TFIIB) (Figure 3.9). Accurate transcription of a gene
by RNA polymerase II requires the assembly of transcription factors at the promoter.
TFIIB localizes in the nucleus with transcription factors IID and IIA where it forms a pre-
initiation complex of RNA polymerase II. TFIID interacts speciﬁcally with the TATA box,
TFIIA with RNA polymerase II and TFIIB functions as a bridge linking the complex
(Deng and Roberts, 2007). It is not known whether Chlamydomonas FAP20 has similar
functions or how it localizes in the flagella or the nucleus. Apart from the flagellar
proteome analysis (Pazour et al., 2005) and Gli transcription factors of the Sonic
hedgehog signaling pathways (Haycraft et al., 2005), transcription factors have not
previously been shown to localize to the flagellum, and if FAP20 functions here it would
be a novel system for transcriptional control.


51
Figure 3.9: Conserved domains of Chlamydomonas FAP20. Fap20 belongs to the
DUF667 superfamily. The FAP20 sequence was blasted for conserved domains on
ncbi.nlm.nih.gov.
Eg5 motor domain was pulled down with GST-IFT172 C-term. This kinesin
belongs to the Kinesin-5 family (BimC family). The rate of bipolar spindle assembly
depends on the MT-gliding velocity of the mitotic kinesin Eg5 in mammals. During
mitosis they have an essential role in pushing the spindle poles to opposite sides of the
cell by pushing the astral MT in opposite directions (Valentine et al., 2006). The N-
terminal region containing the motor domain of Eg5 was the only part of the protein
whose sequence was known when this project started, and therefore I only cloned and
expressed this region to use in the following pull-down experiments. It is intriguing if an
Eg5-like protein interacts with IFT172 and the possible functional implications of such
an interaction are unclear. Nevertheless, we decided to pursue this further.
Figure 3.10: Conserved domains of Chlamydomonas Eg5. Eg5 belongs to the
kinesin-5 family of kinases. The kinesin-5 family has a characteristic BimC box
domain. The Eg5 sequence was blasted for conserved domains on ncbi.nlm.nih.gov.


52
3.8. Construction and purification of MBP-ARFA1A, MBP-FAP20 and
MBP-Eg5 motor domain fusion proteins
To assess if the three chosen proteins, ARFA1A, FAP20 and Eg5 motor domain,
identified in the pull-down analysis indeed interacted with CrEB1 and/or IFT172 C-term,
I set out to test whether the three proteins could bind to either IFT172 or CrEB1 in MBP
pull-down assays. The cDNAs corresponding to the three proteins were first cloned into
the pMalC2 vector (See vector map in appendix D) and transformated into DH10α E.coli
cells. This resulted in fusion of maltose-binding protein (MBP) in-frame to the three
genes and MBP-ARFA1A, MBP-FAP20, and MBP-Eg5 motor domain was constructed.
The sequences were verified for correct insertion. To express the fusion proteins the E.
coli cells were induced with IPTG and the MBP fusion proteins purified on amylose
beads. To check the purity of the MBP-fusion proteins, the beads were run on a SDS-
PAGE gel, and Coomassie stained (Figure 3.11). On the gel there is some, but not a lot
of contamination with E.coli proteins and the apparent molecular weights of the purified
fusion proteins are very close to the theoretical values (See table 3.2). Due to time
constraints, I decided to use these protein preparations for pull-down analysis without
further purification.
3.9. MBP pull-down analysis of ARFA1A, FAP20 and Eg5 motor
domain
To investigate the potential protein-protein interactions, a MBP pull-down experiment, in
which a Chlamydomonas wild type (CC-124) flagella lysate was mixed with either MBP,
MBP-ARFA1A, MBP-FAP20 or MBP-Eg5 motor domain bound to amylose beads, was
performed. The fusion proteins as well as the bound proteins were analyzed by SDS-
PAGE and immunoblotting using antibodies specific for either IFT172, CrEB1 or alpha-
tubulin (Figure 3.12). The results showed no interaction between FAP20/ARFA1A and
IFT172, EB1 or α-tubulin in this assay (Figure 3.12, lanes 3 and 4).


53
Figure 3.11: Coomassie-stained gel of purified MBP-fusion proteins. Fusion
proteins were purified on amylose beads using affinity chromatography as described in
materials and methods. Flow through (FT) was collected to check how successfully the
procedure extracted the MBP fusion proteins from the input.


54
MW
(kDa)
GenBank ID pI Apparent
MW (kDa)
Peptides Amino
acids
MBP 50.8 5.21 ~60
FAP20 22.2 gi: 159468654 9.59 ~72 11 190
ARFA1A 20.6 gi: 159465365 6.43 ~67 2 181
Eg5 40.4 gi: 159475595 7.62 ~90 1 367
Table 3.2. Theoretical values for ArfA1A, FAP20 and Eg5 motor domain. The values
were calculated from the polypeptide sequences using the pI/MW tool on expasy. MBP
(MalE) sequence was obtained on New England Biolabs webpage as an excerpt from
the entire pMalC2 plasmid sequence. The apparent molecular weight was estimated
from a Coomassie stained gel (Figure 3.8). The apparent molecular weights indicated
include the MBP fusion construct. ”Peptides” refer to the number of peptides by which
the proteins were identified in the Chlamydomonas flagellar proteome (Pazour et al.,
2005). Note that the Eg5 fusion protein only contains the N-terminal motor domain.
FAP20 and ARFA1A could have been false positives from the initial pulldown mass
spectrometry experiment, but a number of other factors, can have interfered with this
assay. It is possible that the MBP fusion causes the proteins to misfold thereby not
exposing their binding sites, or the proteins could be lacking post translational
modifications. ARFA1A is a GTPase, so the presence of GTP could have a great effect
of the folding and activity of this protein. Furthermore, the N-terminus of ARF proteins is
known to be critical for their function (Casanova, 2007; Liu et al., 2009) so adding MBP
to the N-terminus of ARFA1A likely interferes with its binding to other proteins.
An interaction of Eg5 motor domain was seen with both IFT172, IFT139 (though
a very weak band) and alpha tubulin (Figure 3.12, lane 5). Repetitions of the analysis
confirmed the association between IFT172 and MBP-Eg5 motor domain (data not
shown). Since MBP-Eg5 contains a motor domain with conserved MT binding sites
(Figure 3.10; multiple sequence alignment in Appendix C) it is not surprising that an
interaction with alpha tubulin is observed.


55
Figure 3.12. Western blot of pull-down analysis of MBP fusion proteins
mixed with wild type (CC-124) flagella extract. Purified MBP fusion
proteins immobilized on amylose resin (see Figure 3.11) were mixed with
flagellar extract and bound proteins analyzed by SDS-PAGE and western
blot with antibodies against flagellar proteins, as indicated. Note that MBP-
Eg5 motor domain co-precipitates with IFT172, IFT139 and alpha tubulin.
It is more surprising that this domain would interact with IFT172, because if IFT172
were bound to the motor domain of Eg5, Eg5 would likely be inactive. It is possible that
IFT172 interacts with Eg5 via microtubules or that Eg5 is inactive when bound to
IFT172. Further experiments are needed to investigate this further. The interaction seen
between Eg5 and IFT139 could be due to the interaction between IFT172 and IFT139,
since IFT139 and IFT172 are both IFT particle proteins, and this could cause the co-
precipitation. To verify the possible binding of Eg5 to IFT172 and tubulin, it would be
useful to clone and sequence the whole Eg5 coding region. My supervisor Lotte
Pedersen recently obtained a full-length Eg5 cDNA clone and sequenced the entire
coding region. To further examine the potential association of Eg5 with tubulin and
IFT172, and potentially the function of Eg5 in Chlamydomonas, it would be highly
relevant to produce an antibody against the C-terminal region of Eg5, which, in contrast
to the motor domain, displays little sequence similarity to other known kinesins (see Eg5
multiple sequence alignment in Appendix C) and therefore would be more appropriate
for antibody production.


56
A specific inhibitor, called Monastrol, is known to act specifically on the motor
domain of human Eg5 (Cochran et al., 2005). Since the motor domain of
Chlamydomonas and human Eg5 are highly conserved (Figure 3.10; multiple sequence
alignment in appendix Appendix C), it is possible that this specific inhibitor will work on
Chlamydomonas Eg5. A few pilot tests have been conducted to see whether Monastrol
has an effect on Chlamydomonas flagellar length or motility, but so far none have been
observed (data not shown). It would be interesting to test the effect of Monastrol on cilia
assembly or disassembly in mammalian cells where Monastrol is known to inhibit Eg5
(Cochran et al., 2005). If an effect is seen, it will provide some clues as to the function of
Eg5 in Chlamydomonas flagella.
3.10. Further testing of the function of ARFA1A
Although no interaction was observed between ARFA1A and IFT172 or EB1 (Figure
3.12) in the MBP pull-down assay, this is a very interesting possible binding partner
given that an interaction between ARF4 and EB1 was observed in parallel studies in
ciliated Retinal pigment epithelial (RPE) cells (see Section 3.7). The interaction was
probably not seen because, as noted above, adding MBP to the N-terminus of ARF
proteins likely inhibits their association with other proteins (Casanova, 2007; Liu et al.,
2009). Chlamydomonas ARFA1A is most closely related to the human ARF members
ARF1 and ARF3-5. A polyclonal antibody raised against the C-terminus of ARF1 of
human origin was purchased, and tested to see if it was able to detect ARFA1A in
Chlamydomonas flagella. Flagella from wild type (CC-124) cells, along with RPE cells in
interphase or after 72 hours of starvation (kindly supplied by fellow student Tue S.
Jørgensen), as well as HeLa cells supplied by the company as a positive control, were
loaded on an SDS-PAGE gel and immunoblotted with the antibody against ARF1
(Figure 3.13). The ARF antibody does not seem capable of detecting ARFA1A in the
Chlamydomonas flagella. However a single band of the appropriate size is seen in
starved RPE cells, but not seen in the RPE cells in interphase. This could indicate that
ARF1 and related proteins are up-regulated during growth arrest, which is typical for
cilia-associated proteins.


57
Figure 3.13: ARF is detected in
RPE cells after 72 hours of
starvation. Western blot of
Chlamydomonas wild type
flagella (wt; CC-124), RPE cells
starved for 72 hours, non-starved
RPE cells, and non-starved HeLa
cells shows that a single band
corresponding to the size of ARF
proteins is seen in starved RPE
cells.
To further test the antibody, to see if it would be able to detect ARF localization in
Chlamydomonas wild type (CC-124) cells, an IFM was conducted, using the antibody
against ARF and a monoclonal anti-acetylated tubulin antibody to stain the flagella
(Figure 3.14). The antibody revealed weak punctate staining along the entire flagellar
length. This distribution is similar to that seen for several IFT components. Due to time
constraints I have not been able to test this possible co-localization further.


58
Figure 3.14: IFM, using a Glutaraldehyde/NP40 fixation method, showing
localization of ArfA1A in C. reinhardtii wild type (CC-124) cells. Cells were grown
at 20°C and subjected to IFM using a polyclonal antibody specific for human ARF
(red). To detect the flagella and basal bodies an antibody specific for acetylated
tubulin (green) was used. The bottom panels show the flagella tip in focus. IFM
showed that ArfA1A (red) localizes with weak punctate staining along the entire
flagellar length. The strong fluorescence of the cell body is due to autofluorescence of
the cells and was also observed when no primary antibodies were added (not shown).
Scale bar 10 µm.
Merge!ARF!AcTub!
No primary antibody!

Master thesis (Mette Lethan)

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