EFFECTS OF FOCAL SEGMENTAL GLOMERULOSCLEROSIS-ASSOCIATED MUTATIONS ON MYOSIN 1E LOCALIZATION AND ACTIVITY
Item Type Dissertation
Authors Karchin, Jing Bi
Rights Attribution-NonCommercial-NoDerivatives 4.0 International
Download date 06/10/2021 03:54:14
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Link to Item http://hdl.handle.net/20.500.12648/2029 EFFECTS OF FOCAL SEGMENTAL
GLOMERULOSCLEROSIS-ASSOCIATED MUTATIONS
ON MYOSIN 1E LOCALIZATION AND ACTIVITY
By
Jing Bi Karchin
A thesis
Presented to the
Department of Cell and Developmental Biology
In partial fulfillment of the requirements
For the degree of Doctor of philosophy
In the
College of Graduate Studies of
State University of New York, Health Science Center
Approved ______
(Sponsor’s signature)
Date ______
Dedications
This thesis is dedicated to three family members,
My maternal grandfather, Xuezhi Bi (1925-2011)
My grandfather by marriage. Joseph Feitler (1926-2015)
And
My mother, Weiying Bi (1959-)
Whose inspiration and support directly made this thesis possible.
ii Acknowledgments
There have been many people in my life who have helped me. I could not have achieved what I have today without all of their support. Thus, I am using this opportunity to express my gratitude to everyone who supported me through the past few years.
Firstly, I would like to express my sincere gratitude to my thesis advisor Dr.
Mira Krendel for her continuous support of my PhD study and thesis research, for her patience, motivation, and immense knowledge. Her guidance helped me in all the time of research, career development, and writing of this thesis. I could not have imagined having a better advisor and mentor for my PhD study.
Besides my advisor, I would like to thank the rest of my thesis committee:
Dr. Patricia Kane, Dr. Jean Sanger, and Dr. David Mitchell for continuing being in my advisory committee from qualify exam; and Dr. Peter Calvert and Dr. Scott
Erdman for joining in my thesis committee; and more importantly, for all their insightful comments and encouragement.
My sincere thanks also go to Dr. Vladimir Sirotkin and his lab members
Robert Carroll and Michael James, for all their time and assistance on the yeast project. Without them, the yeast project will not be finished or published.
I would also like to say thanks to Dr. Stewart Loh for his guidance on protein purification during my rotation at his lab. His effective teaching and patience gave me a solid biochemistry foundation.
iii I thank the labs of Dr. Jeff Amack, Dr. David Prynue, and Dr. Vladimir
Sirotkin for their critical discussion of research directions and manuscript preparations at the joint lab meetings. I also thank my fellow labmates and technicians for all their help and support, and for all the fun we have had in the last four years.
I would like to thank my families, both near and far, for their endless support and encouragement, and for taking the time editing my thesis.
I would like to especially thank my mother, for inspiring me and supporting me without any conditions, and for loving me and believing in me all these years.
I could not have achieved any of these without her.
Last but not the least, I would like to thank my loving and caring husband,
Josh Karchin, for providing me with unfailing support, continuous encouragement and for keeping me going throughout my years of study and through the process of researching and writing this thesis. This accomplishment would not have been possible without him.
Thank you all!
iv Abstract
Effects of Focal Segmental Glomerulosclerosis-Associated Mutations on
Myosin 1e Localization and Activity
Author’s Name: Jing Bi Karchin Sponsor’s Name: Mira Krendel
Our lab has discovered that an actin-dependent molecular motor called
Myosin 1e (Myo1e) is required for maintaining normal morphology and function in vivo of podocytes, a specialized epithelial cell in the kidney. We have found that
Myo1e-null mice develop proteinuria, and mutations in the MYO1E gene, including missense mutations A159P and T119I, and nonsense mutation Y695X, have been identified in focal segmental glomerulosclerosis (FSGS), a primary kidney disease that often leads to end stage renal disease (ESRD). Based on these findings, we have proposed that Myo1e and especially its motor domain, plays a key role in regulating actin cytoskeleton organization in kidney podocytes.
To study Myo1e activity at the junctions, we have used cell culture systems. We confirmed that Myo1e is a component of the podocyte slit diaphragm using glomerular fractionation assay and immune-gold labeling electron microscopy. Disruption of Myo1e motor activity by point mutation
(A159P) completely disrupted Myo1e cellular localization and led to defective actin assembly at nascent cell-cell contacts. Domain mapping experiments in
MDCK cells have suggested that the Myo1e TH2 domain is necessary, but not sufficient for its localization, but addition of the TH1 domain restores its
v localization to junctions. We have also found that the Myo1e SH3 domain interacts with ZO-1, a slit diaphragm and tight junction protein, in in vitro pulldown assays, which might contribute to ZO-1 exchange activity at the junctions.
Another FSGS-associated Myo1e motor domain mutation (T119I) also caused mis-localization of Myo1e in the cultured mouse podocytes, suggesting loss-of- function of the motor domain mutants. We have also shown that ZO-1 is not recruited to the nascent cell-cell contacts at the same time with the Myo1e T119I mutants.
Finally, by using fission yeast as a model system, we have demonstrated that human kidney disease-associated mutations in fission yeast caused defects in yeast growth and endocytosis processes. Interestingly, after analyzing the colocalization patterns between the FSGS-associated Myo1 mutants and
Chaperone Rng3, we have proposed that these two kidney disease-associated mutants likely possess different disease-causing mechanisms.
Above all, we have concluded that Myo1e motor domain plays an important role in its localization and activity in podocyte actin cytoskeleton, which might be the link to the disease mechanism of FSGS at the molecular level.
vi Table of Contents
Abstract ...... v Table of Contents ...... vii List of Figures ...... xiii List of Tables ...... xviii List of Abbreviations ...... xix Chapter 1 Introduction ...... 1 1.1 Relevance of kidney disease ...... 2 1.2 The development of the kidney ...... 3 1.2.1 The organogenesis of the kidney ...... 3 1.2.2 Glomerulogenesis ...... 6 1.2.3 Development of renal filtration barrier ...... 8 1.3 Epithelial tight junctions in the kidney ...... 12 1.4 Functions of non-muscle myosins in the kidney ...... 13 1.4.1 Myosin 1c ...... 14 1.4.2 Myosin 1e ...... 16 1.4.3 Non-muscle myosin IIA ...... 17 1.4.4 Myosin VI ...... 19 1.4.5 Myosin VIIB ...... 21 1.4.6 Myosin IXA ...... 21 1.5 Steroid-resistant nephrotic syndrome and focal segmental glomerulosclerosis...... 23 1.5.1 NPHS1 ...... 24 1.5.2 NPSH2 ...... 25 1.5.3 PLCε1 ...... 27 1.5.4 MYO1E ...... 28 1.5.5 ACTN4 ...... 29 1.5.6 TRPC6 ...... 30 1.5.7 INF2 ...... 31
vii 1.5.8 CD2AP ...... 33 1.5.9 WT-1 ...... 34 1.6 Rationale, hypothesis, and research question ...... 34 1.7 Thesis organization ...... 35 1.8 Attributions ...... 36 1.9 Reference ...... 40 Chapter 2 Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes ...... 66 2.1 Attributions and Acknowledgments ...... 67 2.2 Abstract ...... 68 2.3 Introduction ...... 69 2.4 Materials and Methods ...... 72 2.5 Results ...... 76 2.5.1 Myo1e is a component of the slit diaphragm in the renal glomeruli . 76 2.5.2 Myo1e colocalizes with the slit diaphragm marker ZO-1 in mature and immature glomeruli ...... 77 2.5.3 Myo1e is recruited to cell-cell contacts in cultured podocytes...... 77 2.5.4 Cells lacking functional Myo1e exhibit defects in actin assembly along cell-cell contacts and in contact spreading...... 79 2.5.5 Myo1e localization to cell-cell junctions requires multiple binding motifs...... 80 2.5.6 Myo1e SH3 domain binds to the C-terminal portion of ZO-1...... 81 2.6 Discussion ...... 83 2.7 References ...... 90 Chapter 3 Dissecting the effects of kidney disease-associated Myo1e motor domain mutants on junction dynamics ...... 116 3.1 Attributions and Acknowledgments ...... 117 3.2 Abstract ...... 118 3.3 Introduction ...... 120
viii 3.4 Material and Methods ...... 123 3.5 Results ...... 127 3.5.1 The kidney disease associated Myo1e mutants do not co-localize with ZO-1 at the podocyte junctions...... 127 3.5.2 T119I mutant of Myo1e is not recruited to the junctions during calcium switch assay...... 127 3.5.3 FRAP analysis reveals faster turnover of mCherry-ZO-1 in cells expressing Myo1e mutants...... 128 3.6 Discussion ...... 130 3.7 Reference ...... 133 Chapter 4 Effects of FSGS-associated mutations on the stability and function of myosin-1 in fission yeast ...... 143 4.1 Attributions and acknowledgments ...... 144 4.2 Abstract ...... 145 4.3 Introduction ...... 146 4.4 Materials and Methods ...... 150 4.5 Results ...... 155 4.5.1 Mutations in the motor domain of Myo1, equivalent to the FSGS- associated Myo1e mutations, result in yeast growth defects ...... 155 4.5.2 A181P but not T140I mutation results in the decreased stability of Myo1 in yeast ...... 156 4.5.3 Myo1 motor domain mutations disrupt localization and function of Myo1 in endocytic actin patches ...... 158 4.5.4 Myo1 motor domain mutants co-localize with the eisosome marker Fhn1 ...... 160 4.5.5 Myo1 mutants co-localize with the Cam2 light chain ...... 162 4.5.6 Chaperone Rng3 is recruited to the A181P but not the T140I Myo1 mutant ...... 164 4.6 Discussion ...... 167 4.6.1 FSGS-associated mutations disrupt myosin-1 function ...... 167
ix 4.6.2 Molecular basis for the effects of motor domain mutations on myosin activity and stability...... 167 4.6.3 Motor domain is necessary for myosin-1 localization...... 169 4.6.4 Myo1e motor domain functions in the kidney...... 170 4.7 References ...... 173 Chapter 5 Discussion and Future Directions ...... 209 5.1 Study of Myo1e domain localization and activity at junctions ...... 210 5.2 Model of Myo1e structural organization ...... 213 5.3 Potential role of Myo1e in endocytosis – what have we learned from mouse model studies? ...... 216 5.4 Conclusions ...... 220 5.5 Reference ...... 222 Appendix Chapter 1 Analysis of the interactions between Synaptopodin and myosin 1e ...... 239 AP 1.1 Introduction ...... 240 AP 1.2 Materials and Methods...... 242 AP 1.3 Results ...... 244 AP1.3.1 Localization of Synaptopodin in podocytes ...... 244 AP1.3.2 Generation and expression of FLAG-tagged Synaptopodin fragments ...... 244 AP1.3.3 Synaptopodin fragments interact with the Myo1e SH3 domain 245 AP1.3.4 Synaptopodin does not co-localize with Myo1e in cultured podocytes ...... 246 AP 1.4 Discussion ...... 247 AP 1.5 Reference ...... 249 Appendix Chapter 2 Investigation of the effects of the Myo1e rigor mutant N164A on the dynamics of the actin cytoskeleton ...... 260 AP 2.1 Introduction ...... 261 AP 2.2 Material and Methods ...... 263 AP 2.3 Results ...... 269
x AP 2.3.1 The rigor mutant Myo1e N164A co-localizes with actin at cell-cell junctions ...... 269 AP 2.3.2 The rigor mutant Myo1e N164A co-localizes with ZO-1 at cell-cell junctions...... 270 AP 2.3.3 The rigor mutant Myo1e N164A is recruited to nascent cell-cell contacts with ZO-1 during calcium switch experiments...... 270 AP 2.3.4 The rigor mutant Myo1e N164A does not alter ZO-1 dynamics at the junctions...... 271 AP 2.3.5 Deletion of the TH2 domain from the Myo1e N164A mutant mimics the phenotype of Myo1e-ΔTH2 in the cells...... 271 AP 2.4 Discussion ...... 273 AP 2.4.1 Is Myo1e N164A a rigor mutant? ...... 273 AP 2.4.2 What is the nature of the long protrusions that are induced by the Myo1e N164A mutant? ...... 274 AP 2.4.3 What is the effect of the rigor mutant on the dynamics of the actin cytoskeleton? ...... 275 AP 2.4.4 Is motor activity required for Myo1e function at the junctions? 276 AP 2.5 Reference ...... 278 Appendix Chapter 3 Visualization of cytoskeletal dynamics in podocytes using adenoviral vectors ...... 294 AP 3.1 Attributions and Acknowledgments ...... 295 AP 3.2 Abstract ...... 296 AP 3.3 Introduction ...... 297 AP 3.4 Reagents and instruments ...... 302 AP 3.5 Methods ...... 304 AP 3.6 Results ...... 312 AP 3.6.1 Adenoviral vector infection has high efficiency and low toxicity...... 312 AP 3.6.2 Expression level of recombinant proteins...... 313
xi AP 3.6.3 Using adenovirally encoded cytoskeletal probes to visualize podocyte cytoskeleton...... 314 AP 3.7 Discussion ...... 319 AP 3.8 Reference ...... 322
xii List of Figures
Figure 1.1. Anatomical overview of renal filtration...... 54
Figure 1.2. Schematic cartoon illustrating the glomerular filtration barrier...... 56
Figure 1.3. The development of kidney visceral epithelial cells (podocytes)...... 58
Figure 1.4. Molecular structure of the slit diaphragm...... 60
Figure 1.5. Schematic domain structures of myosin family of molecular motors discussed in the text...... 62
Figure 2.1 Myo1e is a component of slit diaphragm...... 93
Figure 2.2. Myo1e colocalizes with the slit diaphragm marker ZO-1 in mouse glomeruli and in cultured podocytes...... 95
Figure 2.3. Myo1e is recruited to the nascent contacts in cultured podocytes. .. 97
Figure 2.4. Myo1eA159P mutant does not colocalize with actin in the newly formed contacts...... 99
Figure 2.5. Podocytes expressing mutant Myo1e fail to assemble actin filament bundles along cell-cell boundaries...... 101
Figure 2.6. Myo1e localization to the junctions requires multiple domain interactions...... 103
Figure 2.7. Myo1e SH3 domain interacts with the C terminal portion of ZO-1. . 106
Figure 2.8. Model describing how Myo1e may function in cell-cell junctions. ... 108
Figure S2.1. Localization of endogenous Myo1e and ZO-1 after recovery from a calcium switch assay...... 110
Figure S2.2. Localization of Myo1e constructs in MDCK cells...... 112
Figure S2.3. Tail domain of Myo1e localizes to cell-cell contacts in podocytes. 114
xiii Figure 3.1. Kidney disease associated Myo1e mutants do no co-localize with ZO-
1 at the cell-cell junctions...... 135
Figure 3.2. Myo1e is recruited to the nascent cell-cell contact at the same time as
ZO-1 during the junction assembly...... 137
Figure 3.3. Kidney disease-associated Myo1e mutant T119I is not recruited to the nascent cell-cell junction with ZO-1 at the same time...... 139
Figure 3.4. ZO-1 mobile fractions are increased in cells lacking Myo1e or expressing kidney disease-associated Myo1e mutants...... 141
Figure 4.1. Kidney disease-associated mutations in S. pombe Myo1 result in growth defects and decreased protein stability...... 176
Figure 4.2. Motor domain mutations disrupt Myo1 localization to actin patches.
...... 179
Figure 4.3. Motor domain mutations disrupt dynamics of endocytic actin patches visualized with mCherry-tagged fimbrin Fim1...... 181
Figure 4.4. Myo1 motor domain mutants co-localize with the eisosome marker
Fhn1...... 183
Figure 4.5. Myo1 mutants do not co-localize with filamentous actin labeled by
Lifeact-mCherry...... 185
Figure 4.6. Myo1 light chain Cam2 co-localizes with Myo1 mutants and co- localizes with Rng3 in cells expressing A181P but not the T140I Myo1 mutants.
...... 187
xiv Figure 4.7. Chaperone Rng3 is recruited to the A181P but not the T140I Myo1 mutant in eisosomes...... 189
Figure S4.1. The A181P but not T140I mutation reduces the stability of Myo1 in cell extracts before and after cycloheximide treatment, related to Figure 4.1. . 196
Figure S4.2. Motor domain mutations result in re-localization of Myo1 from actin patches to eisosomes, related to Figure 4.2...... 198
Figure S4.3. Recruitment of mutant Myo1 to eisosomes has no effect on the number of eisosomes in cells, related to Figure 4.4...... 200
Figure S4.4. Chaperon Rng3 is recruited to the G308R but not the G483D Myo1 mutant detected using Cam2-mCherry, related to Figure 4.6...... 202
Figure S4.5. Cam2 co-localizes with Myo1 in patches and eisosomes, related to
Figure 4.6A-C’’ and Figure S4.4A-C’’...... 205
Figure S4.6. Chaperone Rng3 is recruited to the G308R but not the G483D Myo1 mutant, related to Figure 4.7...... 207
Figure 5.1. Cartoon showing Myo1e subdomain interactions...... 227
Figure 5.2. Summary of junction localization of Myo1e constructs...... 229
Figure 5.3. Schematic model of Myo1e structural conformation...... 231
AP Figure 1.1. Domain mappings of Synaptopodin (Synpo) and Myosin 1e
(Myo1e)...... 250
AP Figure 1.2. Expression and localization of Synaptopodin in mouse podocytes.
...... 252
AP Figure 1.3. Synaptopodin interacts with Myo1e in pull down assays...... 254
xv AP Figure 1.4. Synaptopodin does not co-localize with full-length Myo1e in differentiated podocytes...... 256
AP Figure 2.1. The Myo1e N164A mutant co-localizes with actin at the plasma membranes and cell protrusions...... 280
AP Figure 2.2. Categories and quantifications of the protrusions in COS-7 cells.
...... 282
AP Figure 2.3. The Myo1e N164A mutant co-localizes with ZO-1 at the cell-cell junctions, similar as the wild type Myo1e...... 284
AP Figure 2.4. The Myo1e N164A mutant is recruited to the nascent cell-cell contact with ZO-1 during contact assembly...... 286
AP Figure 2.5. The Myo1e N164A mutant does not alter the dynamics of junctional ZO-1...... 288
AP Figure 2.6. N164A mutation does not rescue cellular distribution of Myo1e-
ΔTH2...... 290
AP Figure 2.7. Actin cosedimentation assay shows the binding curve of 2 μM
GST-TH2 fusion protein and increasing amount of F-actin...... 292
AP Figure 3.1. Adenoviral infection of podocytes is characterized by high efficiency and low toxicity...... 325
AP Figure 3.2. Western blot analysis of cell lysates prepared from differentiated podocytes infected with different concentrations of the adenoviral vector encoding GFP-Myo1e...... 327
xvi AP Figure 3.3. Localization of fluorescently labeled cytoskeletal markers in differentiated podocytes...... 329
AP Figure 3.4. Visualization of podocyte cytoskeletal dynamics using live-cell imaging...... 331
AP Figure 3.5. Visualization of podocyte cytoskeletal dynamics using multi-color live cell imaging...... 333
xvii List of Tables
Table 1.1 Epithelial cell tight junctions in the kidney...... 64
Table 1.2 Non-muscle myosins in the kidney...... 65
Table S4.1 S. pombe strains used in this study...... 191
Table 5.1: Knockout mouse models leading to proteinuria ...... 233
AP Table 1.1. Synpo-Long Infusion primer design ...... 258
xviii List of Abbreviations
ABM Antibiotic-antimycotic mix
AP Appendix chapter
ATP Adenosine triphosphate
CBD Cargo binding domain
COS-7 Escherichia coli and simian kidney-7
CVL Crude viral lysate
DAD Diaphanous autoregulatory domain
DID Diaphanous inhibitory domain
DMEM Dulbecco’s Modified Eagle Medium
ELC Essential light chain
ENaC Epithelial Na+ channel
ESRD End stage renal disease
FBS Fetal Bovine Serum
FERM Protein 4.1, ezrin, radixin, moesin
FH1/2 Formin homology 1/2
Fig Figure
FP Foot process
FRAP Fluorescence recovery after photobleaching
FSGS Focal segmental glomerulosclerosis
GAP GTPase activating protein motif
GBM Glomerular basement membrane
GTP Guanine triphosphate
HBSS Hank’s Balanced Salt Solution
HEK293 Human embryonic kidney 293 cells
IQ Isoleucine and Glutamine
xix MDCK Madin-Darby canine kidney
MLCK Myosin light chain kinase
MyTH4 Myosin tail homology 4
MTT Methylthiazolyldiphenyl-tetrazolium bromide
NS Nephrotic syndrome
PFU Plaque forming units
RLC Regulatory light chain
RPMI Roswell Park Memorial Institute medium
SH3 Src homology 3
SRNS Steroid resistant nephrotic syndrome
SSNS Steroid sensitive nephrotic syndrome
TH1/2 Tail homology 1/2
TRIM Tripartite motif
TRPC6 Transient receptor potential action channel 6
VEGF Vascular endothelial growth factor
xx Chapter 1 Introduction
Chapter 1
Introduction
1 Chapter 1 Introduction
1.1 Relevance of kidney disease
Kidneys are bean-shaped organs that reside on both sides of the lower back in humans (Fig. 1.1A), with outer darker layer called cortex and inner light layer named medulla. In mammals, kidneys perform both endocrine and exocrine functions. Their excretory activity allows the production of urine, through which metabolic waste products are eliminated (Rhoades and Pflanzer, 1995). The kidneys perform their endocrine function by producing several hormones and enzymes, such as erythropoietin, active form of Vitamin D, and renin (DeLuca,
1973; Santoro et al., 2015; Souma et al., 2015). Additionally, the kidneys also regulate acid-base balance, monitor the concentration of the electrolytes, and regulate the osmolality of the body fluids (Rhoades and Pflanzer, 1995).
The smallest functional unit in the kidney is called nephron (Fig. 1.1B), which is comprised of different renal tubules that are responsible for ion and water reabsorption, and a ball-shaped structure called glomerulus, where the plasma ultrafiltration occurs. The cross section of the glomerulus is shown in Fig.
1.1C. Blood flows into the glomerulus through afferent arteriole (AA), and is filtered into the urinary space (US), and eventually collected by the efferent arteriole (EA). During the filtration process, blood flows through the glomerular filtration barrier (Fig. 1.2), which consists of fenestrated endothelial cells, glomerular basement membrane, and the slit diaphragm between neighboring podocyte foot processes, to allow water, gas, ions and waste product pass through and to retain large sized molecules such as proteins.
2 Chapter 1 Introduction
Focal segmental glomerulosclerosis (FSGS) is a common form of nephrotic syndrome, which can often progress to end stage renal disease
(ESRD). The defining feature of FSGS is massive proteinuria (protein loss from the urine), and often accompanied by hypoalbuminemia and edema (D’Agati et al., 2011). Most forms of proteinuria involve defects in podocytes, specialized epithelial cells residing on the outer surface of glomerular capillary loops.
Podocytes form extensions known as foot processes that interdigitate with those from adjacent cells (Jefferson et al., 2011). Slit diaphragms are cell-cell junctions that bridge adjacent foot processes and are highly permeable to water and small solutes but impermeable to proteins. These unique cell adhesion complexes possess some features of both tight and adherens junctions (Fukasawa et al.,
2009; Reiser et al., 2000). In glomerular disease, podocyte slit diaphragm and the actin cytoskeleton are often targeted by pathological processes (Welsh and
Saleem, 2011).
1.2 The development of the kidney
1.2.1 The organogenesis of the kidney
The kidney, along with other components of the urogenital system, arises from the intermediate mesoderm of the early embryo (Carlson, 2013a; Gilbert,
2014). There are three major stages in the development of the mammalian kidney: pronephros, mesonephros and metanephros. The first stage is also considered the kidney that is only found in the lowest vertebrates, but only the
3 Chapter 1 Introduction last stage will act as a functional kidney in mammals (Carlson, 2013b; Gilbert,
2014; Saxén and Sariola, 1987).
The appearance of the pronephros marks the beginning of the kidney organogenesis. Pronephric duct and pronephric tubules arise separately from the intermediate mesoderm by mesenchyme-epithelial transformation. Eventually, the pronephric tubules and the anterior portion of pronephric duct degenerate, only leaving the posterior portion of the pronephric duct, which serves as part of the excretory system in some mammals and is also known as the Wolffian duct or nephric duct (Kuure et al., 2000). The pronephros in higher vertebrates does not function, but it is required for the induction of the later kidney structures.
The middle portion of the nephric duct induces the adjacent mesenchyme from the intermediate mesoderm to form a new set of the tubules, known as mesonephric tubules, which is the early form of the mesonephros. The mesonephros is also the stage for the development of the aorta and gonads
(Dressler, 2006; Kuure et al., 2000). The nephric duct continues to elongate and attaches to the cloaca, which differentiates upon sex specification (Carlson,
2013b).
As the nephric duct attaches to the cloaca, the cells of the posterior portion of the nephric duct outgrow and form the ureteric bud, which is essential for the formation of the metanephric kidney. As the ureteric bud grows into the posterior portion of the intermediate mesoderm, the cells in the mesoderm that
4 Chapter 1 Introduction surround the ureteric bud form the metanephrogenic blastema (metanephrogenic mesenchyme) (Carlson, 2013b; Uetani and Bouchard, 2009). The ureteric bud receives signals from the metanephrogenic blastema and undergoes subsequent branching. Eventually, the branching of the ureteric bud leads to the formation of the collecting ducts within the kidney (Burrow, 2000). Reciprocally, the ureteric bud also induces the metanephrogenic mesenchyme to accumulate, which leads to the formation of the nephrons, the basic functional units of the kidney.
Vertebrate nephrogenesis is more complex than other organogenesis processes because it includes three kidney forms during development. It is essential to understand the kidney at its embryonic developmental stage, since it is the foundation of developing into mature kidney. Pronephros in human kidney is a temporary organ that is not functional, and usually degenerates or becomes another organ precursor before the development of the mature kidney (Saxén and Sariola, 1987). In lower vertebrates, such as fish and amphibians, the pronephros stage of the kidney is equivalent to the embryonic kidney in humans and mice. They both are induced early during the development, and share similar transcription factor expression patterning, and tissue differentiation events.
Moreover, the pronephros in all fish and amphibians are well developed to a functional organ (Vize et al., 2003). Therefore, researchers have used other model systems, such as Xenopus and zebrafish to study embryonic kidney development and gene expression patterns (Gerlach and Wingert, 2013; Hensey et al., 2002; Jones, 2005; Ryffel, 2003). In addition, in Drosophila, the renal
5 Chapter 1 Introduction structure called “Malpighian tubule” is structurally equivalent to the ureteric bud and functionally similar to the renal tubules in higher vertebrates. Thus, the
Drosophila Malighian tubule structure and function have been explored in the past, and used as a model system to study tubulogenesis in vertebrates (Cagan,
2003; Denholm et al., 2003; Jung et al., 2005).
1.2.2 Glomerulogenesis
The comma-shaped body of the developing glomerulus first appears after the cells from the metanephrogenic blastema receive signals from the branching ureteric bud and form pre-tubular aggregates (renal vesicle) (Burrow, 2000;
Horster et al., 1999). As cells continue to differentiate, an S-shaped body forms.
During this process, the distal end and proximal end of the tubules are identified, as well as the distal and proximal clefts, which lead to the formation of the lumens of renal tubules (Kriz, 2007). The induction signals from the ureteric bud also induce the metanephrogenic mesenchyme to undergo mesenchyme-to- epithelial transition to transform into epithelial cells, which are the precursors of the specialized epithelial cells termed podocytes (Carlson, 2013b), the epithelial cells of the proximal tubules, and the epithelial cells of the distal tubules (Horster et al., 1999; Saxén and Sariola, 1987). At this stage, the migratory endothelial cells receive signals, presumably from podocyte precursor cells, to settle into the distal cleft of the S-shaped body, on the other side of the podocytes precursor cells (Burrow, 2000; Kuure et al., 2000), forming the first capillary loop. As the glomerulus matures, the capillary loop further divides into six to eight loops
6 Chapter 1 Introduction
(Quaggin and Kreidberg, 2008). There are some debates about the origin of these endothelial cells (Saxén and Sariola, 1987). Some have believed that they are also originated from the metanephrogenic mesenchyme, while others have suggested that they are from a different cell fate outside the kidney. The classical experiment that supported the latter evidence was the chimera glomeruli grafting experiment, in which the rodent fetal kidney rudiments were grafted onto quail chorioallantoic membranes. Using immunolabeling of the nuclear marker of the quail cells, the origin of the glomerular endothelial cells in the rodent kidney grafts were verified to be host origin (Abrahamson, 1991; Saxén and Sariola, 1987).
The cells from the comma- and S-shaped body stages will develop into the rest of the nephron components, such as the glomerulus and the renal tubules. After the endothelial cells settle in, the development of the glomerulus enters the capillary loop stage, in which the glomerular basement membrane (GBM) is assembled. Studies have demonstrated that the GBM is originated from a mixed lineage. At the beginning, it appears to be two basement membranes, endothelial basement membrane and early podocytes basement membrane, observed by electron microscopy (Abrahamson, 1991). During the development, the two basement membranes fuse, accompanied by a change in expression of some molecular components of the basement membranes, mainly collagen and laminin
(Dressler, 2006; Kriz, 2007). The similar grafting experiment that mentioned above also provided further information about the composition of the GBM. The evidence suggested that the mature GBM has a dual origin, in which the visceral
7 Chapter 1 Introduction podocytes and endothelial cells both contribute to the GBM biosynthesis and maturation (Abrahamson, 2009, 1991; Horster et al., 1999; Saxén et al., 1986).
1.2.3 Development of renal filtration barrier
The glomerular filtration barrier is comprised of three components: fenestrated endothelial cells, glomerular basement membrane, and the slit diaphragms between the neighboring foot processes of the visceral epithelial cells (podocytes) (Fig. 1.2). The precursor of the structure is first initiated during the S-shaped stage. During the mesenchyme-to-epithelium transition, the metanephrogenic mesenchyme is induced to transform to epithelial-like cells, one of which is the primitive podocytes.
The early podocytes form a layer of columnar epithelial cells with the junctional belt at the apical region (Fig. 1.3A) (Kriz, 2007; Quaggin and Kreidberg,
2008), possibly containing tight junction protein such as ZO-1 and occludins. As the development continues, the podocytes transform to mesenchymal-like cells, but still maintain the epithelial characteristics (Fig. 1.3B-C). During this process, the podocytes become flatter and gradually lose their apical cell attachment. The junctional belt between neighboring podocytes moves from the apical region towards the basolateral region (Kriz, 2007). Nephrin and CD2AP, which are two major components of the podocyte slit diaphragm, are first detected in the capillary loop stage, and eventually become concentrated in the basolateral region and in close proximity to glomerular basement membrane (Li et al., 2000).
8 Chapter 1 Introduction
In addition, podocytes form long protrusions called primary processes, which subdivide into many secondary processes known as foot processes. As the glomerulus matures, these foot processes interdigitate with the ones from neighboring podocytes and entirely cover capillaries. At this stage, podocyte cell bodies completely separate from one another and only interact via neighboring foot processes (Fig. 1.3D). The spaces between neighboring foot processes are called filtration slits, and are covered by a protein complex known as the slit diaphragm, which is one of the major components of the glomerular filtration barrier. It is comprised of transmembrane proteins nephrin, Neph1, podocin, cytoplasmic adaptor proteins ZO-1, CD2AP, and the actin cytoskeleton associated proteins Myo1e and α-actinin-4 (Fig. 1.5). The adaptor proteins and cytoskeleton associated proteins link the actin cytoskeleton to the transmembrane proteins, together maintaining the integrity of the slit diaphragm.
Glomerular endothelial cells originate from outside the kidney and migrate into the distal cleft of the S-shaped body (Abrahamson, 2009). Initially, these endothelial cells are cuboidal shaped and lack fenestrations (Ichimura et al.,
2008; Satchell and Braet, 2009). After entering the capillary loop stage, the endothelial cells change morphology and become thinner in the cytoplasm.
Before this stage, maturing podocytes, located on the other side of the basal membrane, start sending a signal, specifically vascular endothelial growth factor
(VEGF), to the endothelial cells to initiate “pore” formation (Breier et al., 1992;
Esser et al., 1998). The theories of how endothelial cells respond to VEGF and
9 Chapter 1 Introduction form fenestrations are still in debate. One of the theories has suggested the importance of the rearrangement of the actin cytoskeleton. The VEGF secreted from podocytes binds the VEGF receptor on endothelial cells, which in sequence activates the Rac pathway downstream signaling molecule. Rac activation is well known for regulating the actin cytoskeleton, and eventually leads to fenestration formation (Eriksson et al., 2003). An opposing theory on the formation of fenestrations in endothelial cells states that they arise from the preexisting intracellular vesicles (Satchell and Braet, 2009), and then by fusion of these vesicles with both apical and basal membranes (Dvorak et al., 1996). Once the fenestrations are fully developed, they lose their diaphragms or any protein complexes expressed around the pore (Ioannidou et al., 2006). Glomerular endothelial cells also have their own basal membrane, which contributes to the mature glomerular basement membrane formation.
The glomerular basement membrane (GBM) is the third component of the glomerular filtration barrier. It is located between the podocytes and fenestrated endothelial cells. The maturation of the GBM starts from the beginning of glomerulogenesis. The immature podocytes and endothelial cells both secrete their own basal membranes. As maturation proceeds, these two membranes fuse together to form the final state of basement membrane (Saxén et al., 1986).
Collagen I expression is gradually lost from the initial basal membranes before fusion, while Collagen IV becomes the major collagen in the GBM composition.
During this process, other molecular components of the GBM also shift. In the
10 Chapter 1 Introduction comma- and S-shaped body stages, the main components of the GBM are collagen IV α1 and α2 chains (COL4A1 and COL4A2), and laminin-1 (α1β1γ1 heterotrimer). When entering the capillary loop stage, COL4A3, COL4A4, and
COL4A5 are required, in addition to the original two expressed, and laminin-11
(α5β2γ1 heterotrimer) replaces laminin-1. After the final fusion event and towards the maturation of the glomerulus, the mature GBM is comprised of COL4A3,
COL4A4, and COL4A5, and laminin-11. Other than the collagen IV and laminin, agrin and nidogen are also expressed in the GBM (Miner, 1999, Miner, 2012).
Agrin, as a major heparin sulfate proteoglycan in the GBM, contributes to the net negative charge of the GBM, and is thought to repel any negatively charged molecules, such as albumin (Kanwar et al., 2007). Over the years, however, some studies have shown controversial results, that neither podocyte-specific knockout of agrin nor reducing the anionic sites in the GBM alters the permeability of the GBM or causes proteinuria (Harvey et al., 2007; van den
Hoven et al., 2008). The most recent study by Axelsson et al. has shown that the charge effect of the GBM only applies to small sized molecules, but not large molecules (Axelsson et al., 2011). Interestingly, there is a new theory about albumin permeability across the GBM. Hausmann et al. created a mathematical model, in which the albumin filtration across the GBM is due to the electrophoretic effect by the electric potential generated by the filtrates
(Hausmann et al., 2010). However, the exact mechanism of how the GBM facilitates the renal filtration is still under investigation.
11 Chapter 1 Introduction
1.3 Epithelial tight junctions in the kidney
The ultimate function of the kidney is to maintain homeostasis of water and sault in the human body. In order to do that, each segment of the nephron has to properly select the ions or molecules that are allowed to pass through.
Renal epithelial cells and their tight junctions play an essential role in this process.
From the proximal end of the nephron to the distal end, the permeability of the renal tubules changes from “leaky” epithelium to tight epithelium, corresponding to their physiological roles (Denker and Sabath, 2011). Table 1.1 lists the major epithelia in the nephron and their potential physiological functions. As we noted, the podocyte slit diaphragm is the most complex one of them all.
The podocytes slit diaphragm is a specialized junction (Fig. 1.4). Not only does it contain tight junction proteins such as ZO-1, JAM-A (junction adhesion molecule-A) and occludin, it is also a modified adherens junction with P-cadherin and catenins (Reiser et al., 2000, Fukasawa et al., 2009). Characteristically, the slit diaphragm expresses a special set of proteins such as nephrin, Neph1, podocin, and CD2AP, all playing an essential role in maintaining the structural and functional integrity of the renal filtration barrier (Holzman et al., 1999;
Schwarz et al., 2001, Shih et al., 2001, Harita et al., 2008). The glomerular slit diaphragm undergoes dynamic disassembly and reassembly to facilitate renal filtration, which requires the coordination of the underlying cytoskeleton. In podocyte cell bodies and major processes, the dominant filaments are microtubules and intermediate filaments, which play a role in supporting podocyte
12 Chapter 1 Introduction shape (Faul et al., 2007). In the foot processes, the actin-based cytoskeleton consists of actin filaments, non-muscle myosin II, α – actinin 4 and synaptopodin
(See Appendix chapter 1 for more details), which together play essential roles in linking the podocyte slit diaphragm to the glomerular basement membrane via signaling through integrins and other adaptor proteins. Therefore, the podocyte actin cytoskeleton becomes a major target under many pathological events
(Welsh and Saleem, 2011).
1.4 Functions of the non-muscle myosins in the kidney
Myosins are a diverse family of actin-based molecular motors that play important roles in muscle contraction (Herzog et al., 2015), cargo transportation
(Van Den Berg and Hoogenraad, 2012), tension regulation including cell shape change, cell motility and endocytosis (Diz-Munoz et al., 2013; Nambiar et al.,
2009). Thus far, there are more than 20 classes of myosins found in eukaryotes, which have been classified based on the phylogeny of their motor domains
(Richards and Cavalier-Smith, 2005). The motor domains are highly conserved through evolution and bind to actin in an ATP-dependent manner. They exhibit strong actin binding in the ADP-bound state and weak actin binding in the ATP- bound state. The force generated by motor domain ATP hydrolysis is used to carry cargo along the actin filaments. Myosins have one to six IQ (Isoleucine and
Glutamine) motifs that bind to light chain calmodulin or calmodulin-like molecules, playing a role in stabilizing the myosin motor-neck region and amplifying the force for the movement along actin filaments (Tyska and Warshaw, 2002). The tail
13 Chapter 1 Introduction domains of different classes have their own distinct structures and functions, but within one class they likely perform similar or compensatory functions (Sellers,
2000).
The Class II myosins are known as conventional myosins that include both muscle (skeletal and cardiac) and non-muscle myosins (smooth muscle and other non muscle tissue types). This myosin class exists as a dimer in solution with two heavy chains and four light chains (two regulatory light chains and two essential light chains). Phosphorylation of the regulatory light chains by myosin light chain kinase (MLCK) activates the myosin II motor proteins. Upon tail domain self assembly, they form as a tetramer known as the thick filament between actin filaments, initiating muscle contraction (Trybus, 1994; Szent-
Györgyi, 2004). Unconventional myosins refer to those other than Myosin II, while non-muscle myosins refers to any myosins that are not skeletal or cardiac muscle myosins, but it can be conventional myosin (non-muscle myosin IIs). In this section, we are mainly focusing on non-muscle myosins and the overall information is listed in Table 1.2. The schematic domain structures of these myosins are illustrated in Figure 1.5.
1.4.1 Myosin 1c
Myosin 1c (Myo1c), also previously known as Myosin Iβ from bovine adrenal gland and Myr2 from rat kidney (Reizes et al., 1994; Sherr et al., 1993;
Zhu and Ikebe, 1994; Zhu et al., 1996), is a short tailed Class I myosin (Fig. 1.5).
14 Chapter 1 Introduction
Initial studies on Myosin Iβ showed that it was widely expressed in various tissues, and localizes to the cell leading edge (Sherr et al., 1993; Wagner et al.,
1992). In the rat kidney, it is enriched in the proximal tubule microvilli and plays a role in the recovery of the proximal tubule cells from ischemia (Boyd-White et al.,
2001). Subsequent study from Wagner’s group showed that Myo1c was expressed in rat kidney collecting duct cells and enriched at the plasma membrane (Wagner et al., 2005). It has been suggested that Myo1c plays a role in the delivery and insertion of epithelial Na+ channel (ENaC) to the plasma membrane from the cytosol, and activation of the existing channels at the plasma membrane (Wagner et al., 2005). Myo1c, like other class I myosins, has a highly conserved motor domain that generates force upon ATP-hydrolysis. Its motor activity is usually required for its role in vesicular trafficking (Bond et al., 2011).
Myo1c neck domain contains four IQ motifs that bind to light chain calmodulin, and a short tail domain with positive charged residues that binds to phospholipids at the plasma membrane (Gillespie and Cyr, 2004). In the kidney podocytes,
Myo1c was also found to interact with nephrin and Neph1, transmembrane proteins that support the structure of the slit diaphragms, and play a role in targeting them to the slit diaphragms and maintaining slit diaphragm integrity (Arif et al., 2011). To further study Myo1c role in the kidney, the same group of researchers generated Myo1c-null zebrafish and they discovered that knocking down Myo1c in zebrafish using morpholinos resulted in developmental defects in the fish glomeruli (Arif et al., 2013).
15 Chapter 1 Introduction
1.4.2 Myosin 1e
Myosin 1e (Myo1e) is a long tailed class I myosin (Fig. 1.5) that plays a role in cell migration and adhesion, endocytosis, and cell-cell contacts assembly
(Bi et al., 2013; Cheng et al., 2012; Krendel et al., 2007; Ouderkirk and Krendel,
2014). Myo1e is widely expressed in different tissues in vertebrates (Kim et al.,
2006). In the kidney, it is highly enriched in the podocytes and its specialized junction known as slit diaphragms (Bi et al., 2013; Pierchala et al., 2010). A
Myo1e rat homolog, Myr3, has also been shown to localize to adherens junctions in Hela cells (Stöffler et al., 1998), suggesting a common role of class I myosins activity at the cell-cell contacts. The motor domain of Myo1e is highly conserved and binds to actin with low affinity (El Mezgueldi et al., 2002) in an ATP- dependent manner. Myo1e has a single IQ motif in the neck region, which binds to light chain calmodulin, and a long tail domain consisting of three tail homology
(TH) subdomains. The TH1 domain binds to negatively charged phospholipids with high affinity (Feeser et al., 2010). The TH2 domain is rich in proline residues and highly positively charged. Studies from other groups and our own have shown that the TH2 domain contains an ATP-insensitive actin binding site (Jung and Hammer III, 1994). The very C-terminus of Myo1e is a Src homology 3 (SH3) domain, which is responsible for protein-protein interactions with proline-rich containing proteins (Krendel et al., 2007). Myo1e-null mice developed proteinuria and kidney failure with no other significant syndromes, and the ultrastructures from Myo1e-deficient mice, both global and podocyte-specific knock-out, showed
16 Chapter 1 Introduction foot process effacement, loss of slit diaphragms, and thickening of the glomerular basement membrane (Chase et al., 2012; Krendel et al., 2009), which are similar phenotypes that are found in human focal segmental glomerular sclerosis
(FSGS) patients. Mutations in the Myo1e motor domain have been associated with childhood familial FSGS (Al-Hamed et al., 2013; Mele et al., 2011; Sanna-
Cherchi et al., 2011). A recent study using fission yeast as a model system showed that functional motor domain is required for myosin I activity (Bi et al.,
2015). The exact role of Myo1e at the podocyte slit diaphragm is still under investigation.
1.4.3 Non-muscle myosin IIA (NMMIIA)
Non-muscle myosin IIA (NMMIIA) belongs to class II conventional non- sarcomeric myosin (Fig. 1.5) (Sellers, 2000). There are three non-muscle myosin
IIs in vertebrates, NMMIIA, NMMIIB, and NMMIIC, encoded by MYH9, MYH10, and MYH14, respectively, and they share a high sequence and structural homology (Arrondel et al., 2002; D’Apolito et al., 2002; Simons et al., 1991).
NMMIIA is broadly expressed in most cell types in vertebrates, including skeletal muscle cells, and smooth muscle cells (Löfgren et al., 2003; Morano et al., 2000;
Swailes et al., 2006; Yuen et al., 2009). NMMII exists as a dimer of two heavy chains in solution. Structurally, they consist of two heavy chains, two regulatory light chains (RLC), and two essential light chains (ELC). Phosphorylation in RLC mediates dimerization of NMMII dimers, and without this process, the tail of
NMMII folds back to the head, preventing it from associating with others (Vicente-
17 Chapter 1 Introduction
Manzanares et al., 2009). The coiled-coil tail of NMMII self assembles into bipolar filaments, and this anti-parallel hexamer binds to actin filaments through its motor domain and generates contractile force by moving actin filaments
(Vicente-Manzanares et al., 2009). Mutations in the coiled-coil region have been found to affect NMMIIA assembly (Franke et al., 2005). NMMIIs play an essential role in cell migration and cell adhesion (Even-Ram et al., 2007; Swailes et al.,
2006). In the kidney, NMMIIA is expressed in podocytes and proximal tubules. By binding through Dab2 (Disabled-2), a endocytosis associated protein, NMMIIA associates with megalin (a novel albumin receptor found in proximal tubule microvilli) indirectly, regulating megalin endocytosis in proximal tubule cells
(Hosaka et al., 2009). Pharmacologically inhibiting NMMIIA has been shown to delay inflammation infiltration in TNF-induced acute kidney injury and experimental obstructive nephropathy model (Si et al., 2010), suggesting a role of
NMMIIA in regulating immune cell migration and invasion upon inflammatory response in the kidney. By interacting with MG53 (Mitsugumin 53), a muscle specific tripartite motif containing (TRIM) family protein that initiates membrane repair at the damaged site (Cai et al., 2009a, 2009b; Kim et al., 2014), NMMIIA facilitates MG53-containing vesicles traveling to the damaged membrane site, and inhibition of the motor activity of NMMIIA via pharmacological drug compromises MG53-mediated cell membrane repair (Lin et al., 2012). In kidney proximal tubules, MG53-mediated cell membrane repair can potentially prevent kidney from the inflammatory and fibrotic responses, indicating a role of NMMIIA
18 Chapter 1 Introduction in vesicle trafficking in the kidney proximal tubule cells (Duann et al., 2015).
NMMIIA also plays an essential role in the development of kidney. In developing mouse kidney, mesenchyme-specific MYH9 deletion leads to edema in proximal tubule and renal failure (Recuenco et al., 2014). In zebrafish, NMMIIA plays a role in the development of tubules and endothelial cells (Müller et al., 2011).
Genetic studies have identified several mutations in MYH9 gene that are associated with renal dysfunction (Capria et al., 2004; Freedman et al., 2009a;
Freedman et al., 2009b; Kao et al., 2008; Kopp et al., 2008; Rao and
Balakrishnan, 2009), and most of these mutations reside in the NMMIIA motor domain and coiled-coil tail (Zhang et al., 2012), suggesting the underlying mechanism for disease progression is likely due to lack of motor activity or disassembly of the NMMIIA protein. Interestingly, podocyte-specific MYH9 deletion is not sufficient for mice to acquire glomerular disease. The disease only appears in mice upon secondary stimulation, which is consistent to what is seen in some of the human patients with MYH9 risk allele that their renal function only starts to decline upon HIV infection or other stimulus (Johnstone et al., 2011).
1.4.4 Myosin VI
Mammalian Myosin VI was first identified in pigs (Hasson and Mooseker,
1994). It contains a motor domain that binds to actin in an ATP-dependent manner, a 50 amino acid linker region, and one IQ motif that binds to light chain calmodulin (Fig. 1.5) (Hasson and Mooseker, 1994). Its tail domain contains a coiled-coil region that is responsible for dimerization mediated by cargo binding
19 Chapter 1 Introduction
(Yu et al., 2009) and a C-terminal globular cargo binding domain (CBD). Myosin
VI has been confirmed as a processive motor and the only myosin motor that moves towards the pointed (-) end of the actin filament (Nishikawa et al., 2002;
Rock et al., 2001; Wells et al., 1999). There are 3 splice forms found in mammalian tissues resulting from amino acids insertion to the tail domain: the large insert, the small insert, and no insert (Buss et al., 2001). However, only the isoform with the large insertion to the tail domain was exclusively found in the kidney proximal tubule epithelial cells. In murine adult kidneys, Myosin VI localizes to the apical domain of the proximal tubule epithelium and at the base of the actin-rich microvilli (Biemesderfer et al., 2002). In these cells, Myosin VI associates with clathrin, AP-2 (adaptor protein-2) and Dab-2 (Disabled-2), and is involved in clathrin mediated endocytosis or vesicle trafficking (Buss et al., 2001;
Morris et al., 2002). Myosin VI deficient mice developed renal defects including dilated proximal tubule, reduced intake of horseradish peroxidase, and elevated blood pressure (Gotoh et al., 2010), which are consistent with the secondary effects caused by the defective endocytosis pathways. Yang et al. showed that upon hypertension, Myosin VI localization was redistributed from the top to the base of the microvilli in proximal tubule epithelial cells (Yang et al., 2005), which likely resulted in defects in cargo transport and subsequently an altered plasma membrane composition, and explained the defects in proximal tubule physiology.
Due to the structural similarities among microvilli in the kidney proximal tubule, the microvilli in intestinal brush border and the stereocilia in hair cells, mutations
20 Chapter 1 Introduction in Myosin VI gene also link to the histological abnormalities to the intestinal brush border of Snell’s waltzer mice, human deafness disease, and the developmental defects of the stereocillia (Ahmed et al., 2003; Avraham et al., 1997, 1995;
Melchionda et al., 2001; Self et al., 1999).
1.4.5 Myosin VIIB
Myosin VIIB is primarily expressed in the intestine and kidney, where it localizes to the enterocytes or proximal tubule brush borer microvilli, respectively
(Chen et al., 2001). It has a highly conserved motor domain but with a small insertion in the Loop 2, which has been thought to participate in promoting weak
ADP-Pi binding to actin (Henn and De La Cruz, 2005). Myosin VIIB neck region contains 5 IQ motifs, while the tail domain contains MyTH4 (Myosin tail homology
4)-FERM (protein 4.1, ezrin, radixin, moesin) domain repeats (Fig. 1.5), which have been implicated for a role in microtubule binding (Weber et al., 2004), and a novel SH3 domain that binds proline-rich containing proteins (Chen et al., 2001).
It has been shown that the MyTH4-FERM domain in Myosin VIIA tail is involved in cargo binding (Wu et al., 2011). Due to the structural similarity between Myosin
VIIA and VIIB, the MyTH4-FERM repeats in Myosin VIIB tail might also be involved in cargo binding, subsequently playing a role in vesicular trafficking of essential proteins in kidney proximal tubule cells.
1.4.6 Myosin IXA
21 Chapter 1 Introduction
Myosin IXA (also previously known as Myr7 in rat) is the second class IX myosin identified in mammals. It has the highest expression pattern in the brain and testis, with lower expression level in the kidney, adrenal gland, lung, and spleen (Chieregatti et al., 1998). The head domain of Myosin IXA contains a N- terminal extension, a large insertion in the Loop 2 region binding to actin, and an
ATP binding site. There are total of 6 IQ motifs unevenly distributed in the neck region of human Myosin IXA, which bind to light chain calmodulin (Fig. 1.5). The tail domain of Myosin IXA consists of a zinc ion binding region, a Rho-family
GTPase activating protein motif (GAP) that negatively regulates Rho activity, and a software predicted coiled-coil region at the C-terminus (Chieregatti et al., 1998).
It has been shown that unlike other known myosins, Myosin IXA is a single headed processive motor, with a rate-limiting ATP hydrolysis, high affinity to actin and slow dissociation of Mg2+-ATP from actin. The current model of explanation suggested that the large positively charged insertion in the Loop 2 of the motor domain plays a role in this process (Kambara and Ikebe, 2006; Liao et al., 2010;
Nishikawa et al., 2006; Xie, 2010). In the kidney, Myosin IXA is exclusively expressed in the proximal tubule, at the luminal side of the proximal tubule epithelial cells, playing a role in inhibiting megalin (a multi-ligand receptor at the brush boarder of the proximal tubule epithelial cells and binds to albumin) endocytosis. Proteinuria has also been observed in Myosin IXA knockout mice, which is likely due to the defects in proximal tubule cells protein uptake and endocytosis (Thelen et al., 2015).
22 Chapter 1 Introduction
1.5 Steroid-Resistant Nephrotic syndrome and Focal Segmental
Glomerulosclerosis
Nephrotic syndrome (NS) represents heterogeneity of multiple primary renal disorders that are clinically featured as massive proteinuria (protein loss from the urine), hypoalbuminemia (albumin loss from the blood), edema (swelling of the surrounding renal tissues), and/or hyperlipidemia (an abnormally high concentration of fats or lipids in the blood) (Jefferson et al., 2011). It is the most frequent-diagnosed kidney disease that may lead to end stage renal disease
(ESRD). Focal segmental glomerulosclerosis (FSGS) is a common cause of steroid-resistant nephrotic syndrome (SRNS) in children and adults (Gigante et al., 2011; Gipson et al., 2011; Machuca et al., 2009), and because the treatment methods are limited, patients suffering from SRNS usually progress to end stage kidney disease and renal failure early in life. FSGS is a disease of podocytes, which are specialized epithelial cells in renal glomeruli. Podocyte actin cytoskeleton and its associated proteins play key roles in maintaining podocyte shape and regulating the dynamics of the slit diaphragms (Pavenstädt et al.,
2003; Welsh and Saleem, 2011). In the past two decades, mutations in podocyte genes have been identified through various genetic methods that are linked to
FSGS and SRNS. These mutations may be inherited from older generations, or sporadically generated. The mutations found in familial FSGS are inherited following Mendelian ratios, and can be divided into two categories: autosomal recessive mutations, which are typically considered as loss-of-function mutations,
23 Chapter 1 Introduction and clinically the diseases are most commonly found in children and the diseased person only exists in one generation; and autosomal dominant mutations, which are gain-of-function mutations and diseases are late onset and span multiple generations (Pollak, 2014; Rood et al., 2012). In this section, we will briefly introduce some novel podocyte genes that are associated with FSGS or nephrotic syndrome.
1.5.1 NPHS1
More than 15 years ago, Kestila et al. used a positional cloning approach to identify two major mutations in NPHS1 gene that are associated with the congenital nephrotic syndrome of the Finnish type (NPHS1), an autosomal recessive disorder that affects 1 in 8200 births in Finland (Kestilä et al., 1998;
Männikkö et al., 1995; Olsen et al., 1996). The first mutation, also known as Fin- major, reveals a 2-bp (base pair) deletion in the exon 2 that creates a frame shift and results in a stop codon later in exon 2; The other mutation, termed Fin-minor, is a nonsense mutation in the exon 26 (Patrakka et al., 2000). NPHS1 gene is
29-exon long and encodes for a 1241-amino acid transmembrane protein, nephrin. Nephrin is highly expressed in glomerular podocytes, and enriched in the slit diaphragms (Tryggvason, 1999). Nephrin interacts with another transmembrane protein, podocin, together playing a key role in maintaining normal kidney ultrafiltration. Nephrin also interacts with adaptor protein Nck, linking the plasma membrane to the actin cytoskeleton in podocytes (Jones et al.,
2006; Verma et al., 2006). Immunofluorescence staining in kidney sections from
24 Chapter 1 Introduction
Fin-major or Fin-minor patients with antibodies against nephrin reveals negative staining for nephrin, which indicates the lack of nephrin (Patrakka et al., 2000;
Ruotsalainen et al., 2000). Using the stable HEK293 cell line expressing various nephrin mutants, Liu et al. demonstrated that mutant nephrin only accumulated in the ER region, instead of cell surface. Additionally, nephrin distribution was altered in mutant expressing cells, which suggests that the missense mutations in NPHS1 gene are likely to lead to the protein misfolding, ultimately resulting in a defective intracellular transport of nephrin (Liu et al., 2001). Conventional nephrin knockout mice were born at a normal Mendelian ratio but immediately developed massive proteinuria and died within 24 hours. Renal biopsy of these prenatal/neonatal mice showed podocytes foot processes effacement, absence of the slit diaphragm, and the formation of fibrotic tissue (Putaala et al., 2001;
Rantanen et al., 2002). In order to study nephrin function in the adult kidney,
Juhila et al. generated inducible rat nephrin transgenic mice to prevent perinatal lethality of mice lacking nephrin. These transgenic mice lacked endogenous nephrin expression but expressed rat nephrin induced by doxycycline, and developed a severe renal phenotype (Juhila et al., 2010).
1.5.2 NPHS2
Positional cloning also identified another gene, NPHS2, in which mutations have been associated with autosomal recessive steroid-resistant nephrotic syndrome (Boute et al., 2000). In the initial study conducted by Boute et al., the mutations they identified were missense, nonsense, or deletion of one or more
25 Chapter 1 Introduction nucleotides that resulted in frameshift and premature stop codon. One missense mutation nt413G à A on exon 3 (R138Q) accounts for one third of the mutations detected (Boute et al., 2000). Immunostaining with anti-podocin antibody revealed that kidney biopsy from a patient with the R138Q mutation showed negative staining for podocin (Schwarz et al., 2001), which suggested that the
R138Q mutation results in the loss of function of podocin in kidney podocytes, likely due to the misfolding of the protein. Typically as a rule, patients with
NPHS2 mutations show early onset of FSGS or SRNS within the first two decades of life. However, exceptions exist, and several other mutations in the
NPHS2 gene have been identified to associate with late onset of the disease
(Fotouhi et al., 2013; Karle et al., 2002; McKenzie et al., 2007; Tonna et al.,
2008; Tsukaguchi et al., 2002; Mao et al., 2007; Yu et al., 2005). Among these mutations, a R229Q (nt686G à A) polymorphism definitely caught researchers’ attention. Besides the patients tested, R229Q polymorphism was also found in
3% of healthy controls, but the mutation was only found on one allele, and another missense mutation was found on the opposite allele (Karle et al., 2002).
The authors suggest that for patients with late onset FSGS and/or SRNS, besides screening for R229Q polymorphism on NPSH2 gene, other markers should also be tested. NPHS2 encodes for a transmembrane protein, podocin, which is exclusively expressed in kidney podocytes, specifically in slit diaphragm
(Roselli et al., 2002). Research has shown that the C-terminus of podocin interacts with CD2AP and nephrin, and this interaction is essential for maintaining
26 Chapter 1 Introduction a healthy and functional slit diaphragm (Schwarz et al., 2001). To further study the disease mechanism of loss-of-function of podocin in SRNS and/or FSGS, researchers have generated podocin-deficient mice (Mollet et al., 2009; Ratelade et al., 2008; Roselli et al., 2004). These mice lack endogenous podocin and exhibited similar phenotype as patients with NPHS2 mutations. However, constitutively knocking out NPHS2 in mice resulted in early termination of life
(Roselli et al., 2004). Thus, researchers generated podocyte-specific podocin knockout mice. These mice developed FSGS and tubular damage (Mollet et al.,
2009). To further investigate the effects of the disease associated mutation
R138Q, an equivalent mutation (R140Q) was made in knockin mice by gene targeting and homologous recombination (Philippe et al., 2008).
1.5.3 PLCε1
Mutations in PLCε1 were identified to associate with SRNS or FSGS
(Hinkes et al., 2006). Clinical features of children carrying PLCε1 mutations included severe nephrotic syndromes with massive proteinuria and edema.
Kidney biopsies from PLCε1 patients showed signs of FSGS (Boyer et al., 2010).
The PLCε1 gene codes for PLCε1, which belongs to a family of phospholipase.
PLCε1 has been showed to interact with IQGAP and plays a role in the development of glomeruli (Hinkes et al., 2006). Knockdown plce1 in zebrafish caused the disruption of the barrier formation in the pronephric glomeruli and foot process effacement, which is similar as zebrafish NPHS1 or NPHS2 knockdown
(Hinkes et al., 2006). Recently, PLCε1 has been found to induce TRPC6
27 Chapter 1 Introduction activation in podocytes (Kalwa et al., 2015). Surprisingly, however, PLCε1 deficient mice do not exhibit any renal phenotypes (Jefferson and Shankland,
2007).
1.5.4 MYO1E
MYO1E was identified through linkage analysis and homozygosity mapping from a cohort of pediatric patients with nephrotic syndrome (NS) as a new gene that is associated with autosomal recessive childhood NS (Sanna-
Cherchi et al., 2011). Human MYO1E gene encodes a Class I non-muscle,
Myosin 1e (Myo1e). Previous studies have shown that Myo1e is a component of the slit diaphragm and plays a role in maintaining junction integrity (See Chapter
2) (Bi et al., 2013). Direct exome sequencing and next generation sequencing for candidate genes have identified three mutations in the MYO1E gene (Al-Hamed et al., 2013; Mele et al., 2011; Sanna-Cherchi et al., 2011). The Y695X mutation is a nonsense mutation, which creates a truncated protein lacking the tail domain of Myo1e. Both A159P and T119I mutations are missense mutations located in the motor domain of Myo1e, one is adjacent to the Switch I region that binds to actin (A159P), and the other is part of the P-loop (T119I) that involves in nucleotide binding and ATP hydrolysis (Bi et al., 2015). Mutation of either conserved amino acid will disrupt the motor activity of Myo1e. Using fission yeast as a model system, we demonstrated that these motor domain mutations resulted in loss of function of Myo1 in fission yeast, similar as myo1-null fission yeast strains (See Chapter 4) (Bi et al., 2015). Surprisingly, we found that the
28 Chapter 1 Introduction
T140I mutation in yeast (equivalent to T119I in human FSGS patients) disrupts motor domain activity without decreasing protein stability, whereas the A181P mutation (equivalent to A159P in human FSGS patients) is likely to destabilize the protein in addition to disrupt motor domain activity, suggesting two different disease mechanisms in patients with the A159P mutation and the T119I mutation.
Myo1e knockout mice are viable, but develop proteinuria at about 2-3 weeks of age. Ultrastructures show the thickening of the glomerular basement membrane and podocyte foot processes enfacement, similar to FSGS patients (Chase et al.,
2012; Krendel et al., 2009). Myo1e has also been shown to interact with dynamin and synaptojanin-1 through its SH3 domain, suggesting the involvement in endocytosis (Krendel et al., 2007). We have proposed that deficiency in Myo1e, especially lack of motor activity is the main cause of the defects in podocyte endocytosis, which in turn results in slit diaphragm disorganization, and defective podocyte and basement membrane interaction.
1.5.5 ACTN4
The first autosomal dominant form of familial FSGS was caused by mutations in ACTN4 gene (Kaplan et al., 2000). In humans, ACTN4 gene encodes an actin bundling protein, α-actinin 4, which is broadly expressed with its enrichment in kidney podocyte foot processes (Honda et al., 1998)
(Drenckhahn and Franke, 1988). Over the years, there have been more than a dozen mutations found in ACTN4 gene that are associated with familial or idiopathic FSGS (Dai et al., 2009; Šafaríková et al., 2013; Weins et al., 2005).
29 Chapter 1 Introduction
Most of the mutations reside in the highly conserved actin-binding domain of α- actinin 4, resulting in an increased actin binding affinity of the protein (Kaplan et al., 2000; Weins et al., 2007, 2005), which in consequence alters the dynamics of the actin cytoskeleton in podocytes, suggesting a disease causing mechanism.
Mice lacking ACTN4 are viable but born with a lower Mendelian ratio, suggesting that α-actinin 4 deficiency might cause perinatal lethality. Additionally, proteinuria was observed in most of the littermates. There were no consistent differences in podocyte gene expression or glomerular basement membrane thickness, and no other histological abnormalities were observed outside the kidney (Kos et al.,
2003). In humans, heterozygous alleles of ACTN4 gene lead to autosomal dominant form of FSGS. However, heterozygous mice showed no abnormal symptoms. Together with above observations, it has been suggested that α- actinin 4 deficient mice might also exhibit a recessive form of the disease.
1.5.6 TRPC6
Mutations in TRPC6 gene, encoding the cation channel protein transient receptor potential action channel 6 (TRPC6), have been identified to be associated with FSGS (Heeringa et al., 2009; Winn et al., 2005). These heterozygous mutations cause an autosomal dominant form of the disease with late onset disease progression, with one exception of an early onset progression in human patients, at the age of nine (Heeringa et al., 2009). TRPC6 channels are voltage gated Ca2+ channels and broadly expressed in many tissues
(Freichel et al., 2005). In the kidney, TRPC6 has been shown to be a component
30 Chapter 1 Introduction of the podocyte slit diaphragms, playing a key role in regulating Ca2+ signaling through interactions with nephrin and podocin (Reiser et al., 2005).
Overexpression of wild type TRPC6 in cells causes a hyper-activity of Ca2+ influx
(Heeringa et al., 2009). In nephrin knockout mice, TRPC6 expression was significantly increased, leading to a hyper-activity of Ca2+ influx, which is similar to the phenotype caused by most of the disease associated TRPC6 mutations
(Reiser et al., 2005). All mutations found so far are located in the either N- or C- terminus of TRPC6 protein, which might play a role in its interaction with other proteins (Dryer and Reiser, 2010). TRPC6 deficient mice were viable, and showed no histological abnormalities in other renal tissues. There were no significant differences in the mRNA or protein expression level of podocyte genes or other TRPC genes, in blood pressure readings, or albumin content in the urine at the baseline. However, upon induction of renal injury and hypertension,
TRPC6 deficient mice exhibited less proteinuria compared to wild type mice, suggesting that TRPC6 adversely regulates renal function (Eckel et al., 2011).
1.5.7 INF2
Genetic studies including genotyping diseased families, linkage analysis and direct sequencing identified mutations in the INF2 gene that are associated with autosomal dominant FSGS (Brown et al., 2010). The individuals with INF2 mutations develop later onset FSGS and moderate proteinuria, but progress to
ESRD later, similar to patients with autosomal dominant ACTN4 mutations
(Brown et al., 2010). Histological findings showed podocyte foot process
31 Chapter 1 Introduction effacement, as well as unusual actin bundles in foot processes (Brown et al.,
2010). Variable renal phenotypes have also been reported among diseased family members within one single family (Lee et al., 2011). Genetic studies comparing with sporadic cases indicated that mutations in INF2 gene account significantly for the familial FSGS (Barua et al., 2013; Boyer et al., 2011;
Gbadegesin et al., 2012). INF2 encodes inverted formin 2 (INF2), which belongs to a subset of formin family of diaphanous formins. INF2 contains formin homology 1 (FH1) and 2 (FH2) domains that are responsible for actin polymerization and filament elongation (Faix and Grosse, 2006). The N- and C- termini of INF2 contain diaphanous inhibitory domain (DID) and diaphanous autoregulatory domain (DAD)/WASP homology 2 (WH2) domains, respectively, which form an autoinhibitory conformation and is released by binding of GTP- bound Rho (Goode and Eck, 2007). With the discovery of a second actin binding site, INF2 has been shown to both polymerize and depolymerize actin filaments
(Chhabra and Higgs, 2006). All FSGS associated mutations discovered so far are located to the highly conservative residues in the N-terminal DID of INF2. Using a cell culture system to express some of the FSGS associated INF2 mutants, researchers were able to show that disease associated mutations alter the distribution of INF2 and actin filaments in cells (Brown et al., 2010). This suggests that INF2 mutations causing FSGS are likely due to rearrangements of the podocyte actin cytoskeleton. Mutations in INF2 gene are also associated with a neurological disorder called Charcot-Marie-Tooth (CMT) disease (De Rechter
32 Chapter 1 Introduction et al., 2015). However, whether there is a direct linkage between FSGS and CMT is still under investigation.
1.5.8 CD2AP
CD2AP codes for a cytoplasmic adaptor protein, termed CD2-associated protein (CD2AP), which plays important roles in antigen docking, endocytosis, regulation of epithelial junctions (Dustin et al., 1998; Van Duijn et al., 2010;
Welsch et al., 2005). Researchers generated CD2AP deficient mice and discovered that besides the immune system malfunction, these mice also exhibited severe renal defects and died within 7 weeks of age (Shih et al., 1999).
Ultrastructures from 7 day old CD2AP deficient mice showed podocytes foot process effacement, but with normal endothelial cells and glomerular basement membrane (Shih et al., 1999). Using biochemical and microscopic analysis,
CD2AP has been shown to be a component of the podocyte slit diaphragm and to interact with nephrin through its C-terminus (Shih et al., 2001), and might play a role in glomerulogenesis (Li et al., 2000). Interestingly, CD2AP heterozygous mutant mice did not exhibit proteinuria during the course of the 1 year study period, but histological study showed defects in subcellular structures, similar to
CD2AP-null mice and patients with FSGS (Kim et al., 2003). By direct sequencing CD2AP gene in patients with FSGS, researchers have found several mutations that are associated with sporadic FSGS (Gigante et al., 2009; Löwik et al., 2007).
33 Chapter 1 Introduction
1.5.9 WT-1
WT-1 (Wilms’ Tumor associated gene) encodes for WT-1, a tumor suppressor gene that plays a key role in urogenital development (Kreidberg et al.,
1993). Homozygous mutation in WT-1 gene in mice resulted in embryonic lethality, while heterozygous mutation is associated with urogenital malformation, mental retardation and development of Wilms’ Tumor (Kreidberg et al., 1993).
Mutations in the splice site between exon 8 and 9 or complete deletion of one
WT-1 allele are associated with FSGS phenotype with formation of the lesion and diffuse mesangial sclerosis (see review (Morrison et al., 2008)). WT-1 is expressed in kidney podocytes and regulates the expression of podocalyxin
(Palmer et al., 2001). Using WT-1 knockout mice carrying a yeast artificial chromosome containing the human WT-1 locus, researchers were able to rescue the developmental defect in WT-1-null mice. However, reduced WT-1 expression due to the transgene significantly decreased nephrin and podocalyxin expression level in podocytes, which alone could cause glomerular disease (Guo et al.,
2002). A recent genetic study has identified a novel missense mutation that is responsible for autosomal dominant FSGS (Hall et al., 2015). Overexpressing this mutation in cells resulted in apoptosis and defects in focal contact assembly
(Hall et al., 2015).
1.6 Rationale, hypothesis, and research question
34 Chapter 1 Introduction
Our lab has discovered that an actin-dependent molecular motor called
Myo1e is required for maintaining normal podocyte morphology and glomerular function in vivo (Chase et al., 2012). We have found that Myo1e-null mice develop proteinuria (Krendel et al., 2009), and mutations in the MYO1E gene, including missense mutations A159P and T119I, have been identified in FSGS
(Al-Hamed et al., 2013; Mele et al., 2011; Sanna-Cherchi et al., 2011). Based on these findings, we have proposed that Myo1e plays a key role in regulating actin cytoskeleton organization and slit diaphragm integrity in kidney podocytes.
We have previously identified the important roles of Myo1e in podocytes in vivo, which led us to further study Myo1e functional activity in podocytes in culture. The ultimate goal of this project is to clarify the molecular mechanisms linking Myo1e loss to podocyte injury. Based on the main hypothesis and primary outcomes of the project, I have focused on a more narrow research question for my doctoral thesis: what is the significance of Myosin 1e motor domain activity, and how does it affect the podocyte actin cytoskeleton and junctions?
1.7 Thesis organization
In this thesis, I begin with the modification of a junction model to mimic slit diaphragm assembly, which is used frequently in the subsequent chapters. I also identified the initial Myo1e motor and tail domain activity in cells (Chapter 2).
Then I attempted to use a cell culture system to study any defects that are
35 Chapter 1 Introduction caused by Myo1e motor domain mutations (Chapter 3). In the following chapter, I characterized the mutant Myo1e activity using fission yeast as a model system
(Chapter 4). Finally, I conclude with a discussion and future directions for the project (Chapter 5).
Appendix chapters are the side projects that I have done over the years in the lab. The synaptopodin project was from my rotation project (Appendix
Chapter 1), while the N164A project was to characterize a rigor binding mutant of
Myo1e and its activity in the cells (Appendix Chapter 2). The last chapter is adapted from a methods paper on using adenoviral vectors to visualize podocyte actin cytoskeleton (Appendix Chapter 3).
1.8 Attributions
Chapter 1
Chapter 1 is written by myself and edited by my colleagues for grammatical errors. Most of the materials used are referenced at the end of this chapter.
Permission to use the figures from other articles has been obtained from
Nephron Experimental Nephrology, and the New England Journal of Medicine.
Chapter 2
Chapter 2 is adapted from the following article published in 2013:
Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M.,
2013. Myosin 1e is a component of the glomerular slit diaphragm complex that
36 Chapter 1 Introduction regulates actin reorganization during cell-cell contact formation in podocytes. Am.
J. Physiol. Renal Physiol. 305, 532–544.
I designed and performed most of the experiments with the technical help from Chase, SE and Pellenz, CD, including glomerular fractionation, immunofluorescence staining, live-cell imaging, calcium switch experiments and pull down assays. Chase, SE helped generate Myo1e-null mice and Pellenz, CD isolated and subcloned Myo1e-null podocytes from Myo1e-null mice. Dr. Kurihara,
H conducted the electron microscopy expements and prepared the EM figures.
Dr. Fanning, AS supplied MDCK cells for domain mapping expement. Dr. Krendel,
M and I prepared the figures and wrote the manuscript, with input from Dr.
Kurihara and Dr. Fanning. Permission to reuse it as a thesis chapter has been obtained from Amercian Physiological Society.
Chapter 3
Chapter 3 is written by myself and edited by my colleagues for grammatical errors. I designed and performed all the experiments, analyzed all the data, and prepared all the figures. This chapter is not yet ready for publication, since there are still a few more experiments to perform in order to finish the story.
Chapter 4
Chapter 4 is adapted from the following article published in 2015:
37 Chapter 1 Introduction
Bi, J., Carroll, R.T., James, M.L., Ouderkirk, J.L., Krendel, M., Sirotkin, V.,
2015. Effects of FSGS-associated mutations on the stability and function of myosin-1 in fission yeast. Dis. Model. Mech. doi:10.1242/dmm.020214
J.B., R.T.C., M.K., and V.S. designed the experiments. J.B., R.T.C., M.J., and V.S. performed the experiments. J.L.O. prepared the myo1-A181P construct.
J.B. analyzed the data and prepared Figures. J.B., M.K., and V.S. wrote the manuscript with input from R.T.C and M.J. Permission to reuse it as a thesis chapter has been obtained from Disease Models and Mechanisms.
Chapter 5
Chapter 5 is written by myself and edited by my colleague for grammatical errors.
Some of the materials come from my 2014 AHA predoctoral grant proposal
(14PRE20380534). Dr. Krendel and I have discussed some potential directions.
Appendix chapter 1
Appendix chapter 1 is written by myself and edited by my colleagues for grammatical errors. I designed and performed all the experiments and analyzed all the data, and prepared all the figures.
Appendix chapter 2
Appendix chapter 2 is written by myself and edited by my colleagues for grammatical errors. I designed and performed all the experiments and analyzed all the data, and prepared all the figures.
38 Chapter 1 Introduction
Appendix chapter 3
Appendix chapter 3 is adapted from the following article published in 2014:
Bi, J., Pellenz, C.D., Krendel, M., 2014. Visualization of cytoskeletal dynamics in podocytes using adenoviral vectors. Cytoskeleton (Hoboken). 71,
145–156.
Pellenz, CD and I are co-first authors on this publication. Pellenz, CD designed all viral production procedures with modifications from the manufacture suggestions (Agilent Technologies, Santa Clara, CA). I performed all the experiments with the help from Pellenz, CD, and prepared all the figures. Pellenz,
CD, wrote the Materials and Methods section, while I finished writing the rest of the draft. Dr. Krendel, M edited the manuscipt for publication. Permission to reuse it as a thesis chapter has been obtained from © John Wiley & Sons, Inc.
39 Chapter 1 Introduction
1.9 Reference:
Abrahamson, D.R., 2009. Development of kidney glomerular endothelial cells and their role in basement membrane assembly. Organogenesis 5, 275–287. doi:10.4161/org.7577 Abrahamson, D.R., 1991. Glomerulogenesis in the Developing Kidney. Semin. Nephrol. 11, 375–89. Ahmed, Z.M., Morell, R.J., Riazuddin, S., Gropman, A., Shaukat, S., Ahmad, M.M., Mohiddin, S. a, Fananapazir, L., Caruso, R.C., Husnain, T., Khan, S.N., Riazuddin, S., Griffith, A.J., Friedman, T.B., Wilcox, E.R., 2003. Mutations of MYO6 are associated with recessive deafness, DFNB37. Am. J. Hum. Genet. 72, 1315–1322. doi:10.1086/375122 Al-Hamed, M.H., Al-Sabban, E., Al-Mojalli, H., Al-Harbi, N., Faqeih, E., Al Shaya, H., Alhasan, K., Al-Hissi, S., Rajab, M., Edwards, N., Al-Abbad, A., Al-Hassoun, I., Sayer, J. a, Meyer, B.F., 2013. A molecular genetic analysis of childhood nephrotic syndrome in a cohort of Saudi Arabian families. J. Hum. Genet. 1–10. doi:10.1038/jhg.2013.27 Arif, E., Kumari, B., Wagner, M.C., Zhou, W., Holzman, L.B., Nihalani, D., 2013. Myo1c is an unconventional myosin required for zebrafish glomerular development. Kidney Int. 84, 1154–65. doi:10.1038/ki.2013.201 Arif, E., Wagner, M.C., Johnstone, D.B., Wong, H.N., George, B., Pruthi, P. a, Lazzara, M.J., Nihalani, D., 2011. Motor protein Myo1c is a podocyte protein that facilitates the transport of slit diaphragm protein Neph1 to the podocyte membrane. Mol. Cell. Biol. 31, 2134– 2150. doi:10.1128/MCB.05051-11 Arrondel, C., Vodovar, N., Knebelmann, B., Grünfeld, J.-P., Gubler, M.-C., Antignac, C., Heidet, L., 2002. Expression of the nonmuscle myosin heavy chain IIA in the human kidney and screening for MYH9 mutations in Epstein and Fechtner syndromes. J. Am. Soc. Nephrol. 13, 65–74. Avraham, K.B., Hasson, T., Sobe, T., Balsara, B., Testa, J.R., Skvorak, A.B., Morton, C.C., Copeland, N.G., Jenkins, N. a., 1997. Characterization of unconventional MYO6, the human homologue of the gene responsible for deafness in Snell’s waltzer mice. Hum. Mol. Genet. 6, 1225–1231. doi:10.1093/hmg/6.8.1225 Avraham, K.B., Hasson, T., Steel, K.P., Kingsley, D.M., Russell, L.B., Mooseker, M.S., Copeland, N.G., Jenkins, N. a, 1995. The mouse Snell’s waltzer deafness gene encodes an unconventional myosin required for structural integrity of inner ear hair cells. Nat. Genet. 11, 369–375. doi:10.1038/ng1295-369 Axelsson, J., Sverrisson, K., Rippe, a., Fissell, W., Rippe, B., 2011. Reduced diffusion of charge-modified, conformationally intact anionic Ficoll relative to neutral Ficoll across the rat glomerular filtration barrier in vivo. AJP Ren. Physiol. 301, F708–F712. doi:10.1152/ajprenal.00183.2011 Barua, M., Brown, E.J., Charoonratana, V.T., Genovese, G., Sun, H., Pollak, M.R., 2013. Mutations in the INF2 gene account for a significant proportion of familial but not sporadic focal and segmental glomerulosclerosis. Kidney Int. 83, 316–322. doi:10.1038/ki.2012.349 Bi, J., Carroll, R.T., James, M.L., Ouderkirk, J.L., Krendel, M., Sirotkin, V., 2015. Effects of FSGS-associated mutations on the stability and function of myosin-1 in fission yeast. Dis. Model. Mech. doi:10.1242/dmm.020214 Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M., 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am. J. Physiol. Renal Physiol. 305, 532–544. doi:10.1152/ajprenal.00223.2013 Biemesderfer, D., Mentone, S.A., Mooseker, M., Hasson, T., 2002. Expression of myosin VI within the early endocytic pathway in adult and developing proximal tubules. Am. J. Physiol. Renal Physiol. 282, F785–F794. doi:10.1152/ajprenal.00287.2001
40 Chapter 1 Introduction
Bohrer, M.P., Baylis, C., Humes, H.D., Glassock, R.J., Robertson, C.R., Brenner, B.M., 1978. Permselectivity of the glomerular capillary wall. Facilitated filtration of circulating polycations. J. Clin. Invest. 61, 72–78. doi:10.1172/JCI108927 Bond, L.M., Brandstaetter, H., Sellers, J.R., Kendrick-Jones, J., Buss, F., 2011. Myosin motor proteins are involved in the final stages of the secretory pathways. Biochem. Soc. Trans. 39, 1115–1119. doi:10.1042/BST0391115 Boute, N., Gribouval, O., Roselli, S., Benessy, F., Lee, H., Fuchshuber, a, Dahan, K., Gubler, M.C., Niaudet, P., Antignac, C., 2000. NPHS2, encoding the glomerular protein podocin, is mutated in autosomal recessive steroid-resistant nephrotic syndrome. Nat. Genet. 24, 349–354. doi:10.1038/74166 Boyd-White, J., Srirangam, A., Goheen, M.P., Wagner, M.C., 2001. Ischemia disrupts myosin Iβ in renal tubules. Am. J. Physiol. Cell Physiol. 281, C1326–C1335. Boyer, O., Benoit, G., Gribouval, O., Nevo, F., Pawtowski, A., Bilge, I., Bircan, Z., Deschênes, G., Guay-Woodford, L.M., Hall, M., Macher, M.-A., Soulami, K., Stefanidis, C.J., Weiss, R., Loirat, C., Gubler, M.-C., Antignac, C., 2010. Mutational analysis of the PLCE1 gene in steroid resistant nephrotic syndrome. J. Med. Genet. 47, 445–452. doi:10.1136/jmg.2009.076166 Boyer, O., Benoit, G., Gribouval, O., Nevo, F., Tête, M.-J., Dantal, J., Gilbert- Dussardier, B., Touchard, G., Karras, A., Presne, C., Grunfeld, J.-P., Legendre, C., Joly, D., Rieu, P., Mohsin, N., Hannedouche, T., Moal, V., Gubler, M.-C., Broutin, I., Mollet, G., Antignac, C., 2011. Mutations in INF2 are a major cause of autosomal dominant focal segmental glomerulosclerosis. J. Am. Soc. Nephrol. 22, 239–245. doi:10.1681/ASN.2010050518 Breier, G., Albrecht, U., Sterrer, S., Risau, W., 1992. Expression of vascular endothelial growth factor during embryonic angiogenesis and endothelial cell differentiation. Development 114, 521–532. Brown, E.J., Schlöndorff, J.S., Becker, D.J., Uscinski, A.L., Higgs, H.N., Henderson, J.M., Pollak, M.R., 2010. Mutations in the formin protein INF2 cause focal segmental glomerulosclerosis. Nat. Genet. 42, 72–76. doi:10.1038/ng.505.Mutations Burrow, C.R., 2000. Regulatory molecules in kidney development. Pediatr. Nephrol. 14, 240–253. doi:10.1007/s004670050049 Buss, F., Arden, S.D., Lindsay, M., Luzio, J.P., Kendrick-Jones, J., 2001. Myosin VI isoform localized to clathrin-coated vesicles with a role in clathrin-mediated endocytosis. EMBO J. 20, 3676–3684. doi:10.1093/emboj/20.14.3676 Cagan, R., 2003. The signals that drive kidney development: a view from the fly eye. Curr. Opin. Nephrol. Hypertens. 12, 11–17. doi:10.1097/00041552-200301000-00003 Cai, C., Masumiya, H., Weisleder, N., Matsuda, N., Nishi, M., Hwang, M., Ko, J., Lin, P., Thornton, A., Zhao, X., Pan, Z., Komazaki, S., Brotto, M., Takeshima, H., Ma, J., 2009a. MG53 nucleates assembly of cell membrane repair machinery. Nat. Cell Biol. 11, 56–64. doi:10.1038/ncb1812.MG53 Cai, C., Masumiya, H., Weisleder, N., Pan, Z., Nishi, M., Komazaki, S., Takeshima, H., Ma, J., 2009b. MG53 regulates membrane budding and exocytosis in muscle cells. J. Biol. Chem. 284, 3314–3322. doi:10.1074/jbc.M808866200 Capria, M., Andreucci, M., Fuiano, L., Mancuso, D., Cianfrone, P., Comi, N., Mazza, G., Caglioti, A., Fuiano, G., 2004. Double nucleotidic mutation of the MYH9 gene in a young patient with end-stage renal disease. Nephrol. Dial. Transplant 19, 249–251. doi:10.1093/ndt/gfg475 Carlson, B.M., 2013a. Chapter 6: Establishment of the Basic Embryonic Body Plan, in: Human Embryology and Developmental Biology. pp. 103–104. Carlson, B.M., 2013b. Chapter 16: Urogenital System, in: Human Embryology and Developmental Biology. pp. 376–383. Chase, S.E., Encina, C. V, Stolzenburg, L.R., Tatum, A.H., Holzman, L.B., Krendel, M., 2012. Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am. J. Physiol. Renal Physiol. 303, F1099–106. doi:10.1152/ajprenal.00251.2012
41 Chapter 1 Introduction
Chen, Z.Y., Hasson, T., Zhang, D.S., Schwender, B.J., Derfler, B.H., Mooseker, M.S., Corey, D.P., 2001. Myosin-VIIb, a novel unconventional myosin, is a constituent of microvilli in transporting epithelia. Genomics 72, 285–296. doi:10.1006/geno.2000.6456 Cheng, J., Grassart, A., Drubin, D.G., 2012. Myosin 1E coordinates actin assembly and cargo trafficking during clathrin-mediated endocytosis. J. Cell Sci. Chhabra, E.S., Higgs, H.N., 2006. INF2 is a WASP homology 2 motif-containing formin that severs actin filaments and accelerates both polymerization and depolymerization. J. Biol. Chem. 281, 26754–26767. doi:10.1074/jbc.M604666200 Chieregatti, E., Gärtner, a, Stöffler, H.E., Bähler, M., 1998. Myr 7 is a novel myosin IX- RhoGAP expressed in rat brain. J. Cell Sci. 111 ( Pt 2, 3597–3608. D’Agati, V.D., Kaskel, F.J., Falk, R.J., 2011. Focal segmental glomerulosclerosis. N. Engl. J. Med. 365, 2398–411. doi:10.1056/NEJMra1106556 D’Apolito, M., Guarnieri, V., Boncristiano, M., Zelante, L., Savoia, A., 2002. Cloning of the murine non-muscle myosin heavy chain IIA gene ortholog of human MYH9 responsible for May-Hegglin, Sebastian, Fechtner, and Epstein syndromes. Gene 286, 215–222. doi:10.1016/S0378-1119(02)00455-9 Dai, S., Wang, Z., Pan, X., Chen, X., Wang, W., Ren, H., Feng, Q., He, J.C., Han, B., Chen, N., 2009. ACTN4 gene mutations and single nucleotide polymorphisms in idiopathic focal segmental glomerulosclerosis. Nephron - Clin. Pract. 111, c87–c94. doi:10.1159/000191198 De Rechter, S., De Waele, L., Levtchenko, E., Mekahli, D., 2015. Charcot–Marie– Tooth: Are you testing for proteinuria? Eur. J. Paediatr. Neurol. 19, 1–5. doi:10.1016/j.ejpn.2014.08.004 DeLuca, H.F., 1973. The kidney as an endocrine organ for the production of 1,25- Dihydroxyvitamin D, a calcium-mobilizing hormone. N. Engl. J. Med. 289, 359–365. Denholm, B., Sudarsan, V., Pasalodos-Sanchez, S., Artero, R., Lawrence, P., Maddrell, S., Baylies, M., Skaer, H., 2003. Dual origin of the renal tubules in Drosophia: Mesodermal cells integrate and polarize to establish secretory function. Curr. Biol. 13, 1052– 1057. doi:10.1016/S Denker, B.M., Sabath, E., 2011. The biology of epithelial cell tight junctions in the kidney. J. Am. Soc. Nephrol. 22, 622–625. doi:10.1681/ASN.2010090922 Diz-Munoz, A., Fletcher, D.A., Weiner, O.D., 2013. Use the force: Membrane tension as an organizaer of cell shape and motility. Trends Cell Biol. 23, 47–53. doi:10.1016/j.biotechadv.2011.08.021.Secreted Drenckhahn, D., Franke, R.P., 1988. Ultrastructural organization of contractile and cytoskeletal proteins in glomerular podocytes of chicken, rat, and man. Lab. Invest. Dressler, G.R., 2006. The cellular basis of kidney development. Annu. Rev. Cell Dev. Biol. 22, 509–29. doi:10.1146/annurev.cellbio.22.010305.104340 Dryer, S.E., Reiser, J., 2010. TRPC6 channels and their binding partners in podocytes: role in glomerular filtration and pathophysiology. Am. J. Physiol. Renal Physiol. 299, F689–F701. doi:10.1152/ajprenal.00298.2010 Duann, P., Li, H., Lin, P., Tan, T., Wang, Z., Chen, K., Zhou, X., Gumpper, K., Zhu, H., Ludwig, T., Mohler, P.J., Rovin, B., Abraham, W.T., Zeng, C., Ma, J., 2015. MG53-mediated cell membrane repair protects against acute kidney injury. Sci. Transl. Med. 7, 36–47. Dustin, M.L., Olszowy, M.W., Holdorf, A.D., Li, J., Bromley, S., Desai, N., Widder, P., Rosenberger, F., Van Der Merwe, P.A., Allen, P.M., Shaw, A.S., 1998. A novel adaptor protein orchestrates receptor patterning and cytoskeletal polarity in T-cell contacts. Cell 94, 667–677. doi:10.1016/S0092-8674(00)81608-6 Dvorak, a M., Kohn, S., Morgan, E.S., Fox, P., Nagy, J. a, Dvorak, H.F., 1996. The vesiculo-vacuolar organelle (VVO): a distinct endothelial cell structure that provides a transcellular pathway for macromolecular extravasation. J. Leukoc. Biol. 59, 100–115. Eckel, J., Lavin, P.J., Finch, E. a, Mukerji, N., Burch, J., Gbadegesin, R., Wu, G., Bowling, B., Byrd, A., Hall, G., Sparks, M., Zhang, Z.S., Homstad, A., Barisoni, L.,
42 Chapter 1 Introduction
Birbaumer, L., Rosenberg, P., Winn, M.P., 2011. TRPC6 enhances angiotensin II-induced albuminuria. J. Am. Soc. Nephrol. 22, 526–535. doi:10.1681/ASN.2010050522 El Mezgueldi, M., Tang, N., Rosenfeld, S.S., Ostap, E.M., 2002. The kinetic mechanism of Myo1e (human myosin-IC). J. Biol. Chem. 277, 21514–21. doi:10.1074/jbc.M200713200 Eriksson, A., Cao, R., Roy, J., Tritsaris, K., Wahlestedt, C., Dissing, S., Thyberg, J., Cao, Y., 2003. Small GTP-binding protein Rac is an essential mediator of vascular endothelial growth factor-induced endothelial fenestrations and vascular permeability. Circulation 107, 1532– 1538. doi:10.1161/01.CIR.0000055324.34758.32 Esser, S., Wolburg, K., Wolburg, H., Breier, G., Kurzchalia, T., Risau, W., 1998. Vascular endothelial growth factor induces endothelial fenestrations in vitro. J. Cell Biol. 140, 947–959. Even-Ram, S., Doyle, A.D., Conti, M.A., Matsumoto, K., Adelstein, R.S., Yamada, K.M., 2007. Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat. Cell Biol. 9, 299–309. doi:10.1038/ncb1540 Faix, J., Grosse, R., 2006. Staying in Shape with Formins. Dev. Cell 10, 693–706. doi:10.1016/j.devcel.2006.05.001 Faul, C., Asanuma, K., Yanagida-Asanuma, E., Kim, K., Mundel, P., 2007. Actin up: regulation of podocyte structure and function by components of the actin cytoskeleton. Trends Cell Biol. 17, 428–37. doi:10.1016/j.tcb.2007.06.006 Feeser, E. a, Ignacio, C.M.G., Krendel, M., Ostap, E.M., 2010. Myo1e binds anionic phospholipids with high affinity. Biochemistry 49, 9353–60. doi:10.1021/bi1012657 Fotouhi, N., Ardalan, M., Bonyadi, M.J., Abdolmohammadi, R., Kamalifar, A., Nasri, H., Einollahi, B., 2013. R229Q Polymorphism of NPHS2 Gene in Patients With Late-Onset Steroid-Resistance Nephrotic Syndrome A Preliminary Study. Iran. J. Kidney Dis. 7, 339–403. Franke, J.D., Dong, F., Rickoll, W.L., Kelley, M.J., Kiehart, D.P., 2005. Rod mutations associated with MHY9-related disorders disrupt nonmuscle myosin-IIA assembly. Blood 105, 161–169. doi:10.1182/blood-2004-06-2067 Freedman, B.I., Hicks, P.J., Bostrom, M. a, Cunningham, M.E., Liu, Y., Divers, J., Kopp, J.B., Winkler, C. a, Nelson, G.W., Langefeld, C.D., Bowden, D.W., 2009a. Polymorphisms in the non-muscle myosin heavy chain 9 gene (MYH9) are strongly associated with end-stage renal disease historically attributed to hypertension in African Americans. Kidney Int. 75, 736–745. doi:10.1038/ki.2008.701 Freedman, B.I., Hicks, P.J., Bostrom, M. a., Comeau, M.E., Divers, J., Bleyer, A.J., Kopp, J.B., Winkler, C. a., Nelson, G.W., Langefeld, C.D., Bowden, D.W., 2009b. Non-muscle myosin heavy chain 9 gene MYH9 associations in African Americans with clinically diagnosed type 2 diabetes mellitus-associated ESRD. Nephrol. Dial. Transplant. 24, 3366–3371. doi:10.1093/ndt/gfp316 Freichel, M., Vennekens, R., Olausson, J., Stolz, S., Philipp, S.E., Weissgerber, P., Flockerzi, V., 2005. Functional role of TRPC proteins in native systems: implications from knockout and knock-down studies. J. Physiol. 567, 59–66. doi:10.1113/jphysiol.2005.092999 Fukasawa, H., Bornheimer, S., Kudlicka, K., Farquhar, M.G., 2009. Slit diaphragms contain tight junction proteins. J. Am. Soc. Nephrol. 20, 1491–503. doi:10.1681/ASN.2008101117 Gbadegesin, R. a, Lavin, P.J., Hall, G., Bartkowiak, B., Homstad, A., Jiang, R., Wu, G., Byrd, A., Lynn, K., Wolfish, N., Ottati, C., Stevens, P., Howell, D., Conlon, P., Winn, M.P., 2012. Inverted formin 2 mutations with variable expression in patients with sporadic and hereditary focal and segmental glomerulosclerosis. Kidney Int. 81, 94–99. doi:10.1038/ki.2011.297 Gerlach, G.F., Wingert, R. a, 2013. Kidney organogenesis in the zebrafish: insights into vertebrate nephrogenesis and regeneration. Wiley Interdiscip. Rev. Dev. Biol. 2, 559–85. doi:10.1002/wdev.92
43 Chapter 1 Introduction
Gigante, M., Piemontese, M., Gesualdo, L., Iolascon, A., Aucella, F., 2011. Molecular and genetic basis of inherited nephrotic syndrome. Int. J. Nephrol. 2011, 1–15. doi:10.4061/2011/792195 Gigante, M., Pontrelli, P., Montemurno, E., Roca, L., Aucella, F., Penza, R., Caridi, G., Ranieri, E., Ghiggeri, G.M., Gesualdo, L., 2009. CD2AP mutations are associated with sporadic nephrotic syndrome and focal segmental glomerulosclerosis (FSGS). Nephrol. Dial. Transplant. 24, 1858–1864. doi:10.1093/ndt/gfn712 Gilbert, S.F., 2014. Chapter 12: Paraxial and Intermediate Mesoderm, in: Developmental Biology. pp. 436–447. Gillespie, P.G., Cyr, J.L., 2004. Myosin-1c, the hair cell’s adaptation motor. Annu. Rev. Physiol. 66, 521–545. doi:10.1146/annurev.physiol.66.032102.112842 Gipson, D.S., Trachtman, H., Kaskel, F.J., Greene, T.H., Radeva, M.K., Gassman, J.J., Moxey-Mims, M.M., Hogg, R.J., Watkins, S.L., Fine, R.N., Hogan, S.L., Middleton, J.P., Vehaskari, V.M., Flynn, P. a, Powell, L.M., Vento, S.M., McMahan, J.L., Siegel, N., D’Agati, V.D., Friedman, A.L., 2011. Clinical trial of focal segmental glomerulosclerosis in children and young adults. Kidney Int. 80, 868–878. doi:10.1038/ki.2011.195 Goode, B.L., Eck, M.J., 2007. Mechanism and function of formins in the control of actin assembly. Annu. Rev. Biochem. 76, 593–627. doi:10.1146/annurev.biochem.75.103004.142647 Gotoh, N., Yan, Q., Du, Z., Biemesderfer, D., Kashgarian, M., Mooseker, M.S., Wang, W., 2010. Altered renal proximal tubular endocytosis and histology in mice lacking myosin-VI. Cytoskeleton (Hoboken). 67, 178–192. doi:10.1016/j.biotechadv.2011.08.021.Secreted Guo, J.-K., Menke, A.L., Gubler, M.-C., Clarke, A.R., Harrison, D., Hammes, A., Hastie, N.D., Schedl, A., 2002. WT1 is a key regulator of podocyte function: reduced expression levels cause crescentic glomerulonephritis and mesangial sclerosis. Hum. Mol. Genet. 11, 651– 659. Hall, G., Gbadegesin, R. a., Lavin, P., Wu, G., Liu, Y., Oh, E.C., Wang, L., Spurney, R.F., Eckel, J., Lindsey, T., Homstad, a., Malone, a. F., Phelan, P.J., Shaw, a., Howell, D.N., Conlon, P.J., Katsanis, N., Winn, M.P., 2015. A Novel Missense Mutation of Wilms’ Tumor 1 Causes Autosomal Dominant FSGS. J. Am. Soc. Nephrol. 26, 831–843. doi:10.1681/ASN.2013101053 Harita, Y., Kurihara, H., Kosako, H., Tezuka, T., Sekine, T., Igarashi, T., Hattori, S., 2008. Neph1, a component of the kidney slit diaphragm, is tyrosine-phosphorylated by the Src family tyrosine kinase and modulates intracellular signaling by binding to Grb2. J. Biol. Chem. 283, 9177–9186. doi:10.1074/jbc.M707247200 Harvey, S.J., Jarad, G., Cunningham, J., Rops, A.L., van der Vlag, J., Berden, J.H., Moeller, M.J., Holzman, L.B., Burgess, R.W., Miner, J.H., 2007. Disruption of glomerular basement membrane charge through podocyte-specific mutation of agrin does not alter glomerular permselectivity. Am. J. Pathol. 171, 139–152. doi:10.2353/ajpath.2007.061116 Hasson, T., Mooseker, M.S., 1994. Porcine myosin-VI: Characterization of a new mammalian unconventional myosin. J. Cell Biol. 127, 425–440. doi:10.1083/jcb.127.2.425 Hausmann, R., Kuppe, C., Egger, H., Schweda, F., Knecht, V., Elger, M., Menzel, S., Somers, D., Braun, G., Fuss, A., Uhlig, S., Kriz, W., Tanner, G., Floege, J., Moeller, M.J., 2010. Electrical forces determine glomerular permeability. J. Am. Soc. Nephrol. 21, 2053–2058. doi:10.1681/ASN.2010030303 Heeringa, S.F., Moller, C.C., Du, J., Yue, L., Hinkes, B., Chernin, G., Vlangos, C.N., Hoyer, P.F., Reiser, J., Hildebrandt, F., 2009. A Novel TRPC6 Mutation That Causes Childhood FSGS. PLoS One 4, 1–8. doi:10.1371/Citation Henn, A., De La Cruz, E.M., 2005. Vertebrate myosin VIIb is a high duty ratio motor adapted for generating and maintaining tension. J. Biol. Chem. 280, 39665–39676. doi:10.1074/jbc.M507667200 Hensey, C., Dolan, V., Brady, H.R., 2002. The Xenopus pronephros as a model system for the study of kidney development and pathophysiology. Nephrol. Dial. Transplant 17 Suppl 9, 73–74.
44 Chapter 1 Introduction
Herzog, W., Powers, K., Johnston, K., Duvall, M., 2015. A new paradigm for muscle contraction. Front. Physiol. 6, 1–11. doi:10.3389/fphys.2015.00174 Hinkes, B., Wiggins, R.C., Gbadegesin, R., Vlangos, C.N., Seelow, D., Nürnberg, G., Garg, P., Verma, R., Chaib, H., Hoskins, B.E., Ashraf, S., Becker, C., Hennies, H.C., Goyal, M., Wharram, B.L., Schachter, A.D., Mudumana, S., Drummond, I., Kerjaschki, D., Waldherr, R., Dietrich, A., Ozaltin, F., Bakkaloglu, A., Cleper, R., Basel-Vanagaite, L., Pohl, M., Griebel, M., Tsygin, A.N., Soylu, A., Müller, D., Sorli, C.S., Bunney, T.D., Katan, M., Liu, J., Attanasio, M., O’toole, J.F., Hasselbacher, K., Mucha, B., Otto, E. a, Airik, R., Kispert, A., Kelley, G.G., Smrcka, A. V, Gudermann, T., Holzman, L.B., Nürnberg, P., Hildebrandt, F., 2006. Positional cloning uncovers mutations in PLCE1 responsible for a nephrotic syndrome variant that may be reversible. Nat. Genet. 38, 1397–1405. doi:10.1038/ng1918 Holzman, L.B., John, P.L.S.T., Kovari, I. a., Verma, R., Holthofer, H., Abrahamson, D.R., 1999. Nephrin localizes to the slit pore of the glomerular epithelial cell: Rapid communication. Kidney Int. 56, 1481–1491. doi:10.1046/j.1523-1755.1999.00719.x Honda, K., Yamada, T., Endo, R., Ino, Y., Gotoh, M., Tsuda, H., Yamada, Y., Chiba, H., Hirohashi, S., 1998. Actinin-4, a Novel Actin-bundling Protein Associated with Cell Motility and Cancer Invasion. J. Cell Biol. 140, 1383–1393. Horster, M.F., Braun, G.S., Huber, S.M., 1999. Embryonic renal epithelia: induction, nephrogenesis, and cell differentiation. Physiol. Rev. 79, 1157–1191. Hosaka, K., Takeda, T., Iino, N., Hosojima, M., Sato, H., Kaseda, R., Yamamoto, K., Kobayashi, A., Gejyo, F., Saito, A., 2009. Megalin and nonmuscle myosin heavy chain IIA interact with the adaptor protein Disabled-2 in proximal tubule cells. Kidney Int. 75, 1308–1315. doi:10.1038/ki.2009.85 Ichimura, K., Stan, R. V, Kurihara, H., Sakai, T., 2008. Glomerular endothelial cells form diaphragms during development and pathologic conditions. J. Am. Soc. Nephrol. 19, 1463– 1471. doi:10.1681/ASN.2007101138 Ioannidou, S., Deinhardt, K., Miotla, J., Bradley, J., Cheung, E., Samuelsson, S., Ng, Y.-S., Shima, D.T., 2006. An in vitro assay reveals a role for the diaphragm protein PV-1 in endothelial fenestra morphogenesis. Proc. Natl. Acad. Sci. U. S. A. 103, 16770–16775. doi:10.1073/pnas.0603501103 Jefferson, J.A., Nelson, P.J., Najafian, B., Shankland, S.J., 2011. Podocyte Disorders: Core Curriculum 2011. Am. J. Kidney Dis. 58, 666–677. doi:10.1053/j.ajkd.2011.05.032 Jefferson, J.A., Shankland, S.J., 2007. Familial nephrotic syndrome: PLCE1 enters the fray. Nephrol. Dial. Transplant. 22, 1849–1852. doi:10.1093/ndt/gfm098 Johnstone, D.B., Zhang, J., George, B., Léon, C., Gachet, C., Wong, H., Parekh, R., Holzman, L.B., 2011. Podocyte-specific deletion of Myh9 encoding nonmuscle myosin heavy chain 2A predisposes mice to glomerulopathy. Mol. Cell. Biol. 31, 2162–2170. doi:10.1128/MCB.05234-11 Jones, E. a, 2005. Xenopus: a prince among models for pronephric kidney development. J. Am. Soc. Nephrol. 16, 313–321. doi:10.1681/ASN.2004070617 Jones, N., Blasutig, I.M., Eremina, V., Ruston, J.M., Bladt, F., Li, H., Huang, H., Larose, L., Li, S.S.-C., Takano, T., Quaggin, S.E., Pawson, T., 2006. Nck adaptor proteins link nephrin to the actin cytoskeleton of kidney podocytes. Nature 440, 818–823. doi:10.1038/nature04662 Juhila, J., Lassila, M., Roozendaal, R., Lehtonen, E., Messing, M., Langer, B., Kerjaschki, D., Verbeek, J.S., Holthofer, H., 2010. Inducible nephrin transgene expression in podocytes rescues nephrin-deficient mice from perinatal death. Am. J. Pathol. 176, 51–63. doi:10.2353/ajpath.2010.080843 Jung, A.C., Denholm, B., Skaer, H., Affolter, M., 2005. Renal tubule development in Drosophila: a closer look at the cellular level. J. Am. Soc. Nephrol. 16, 322–328. doi:10.1681/ASN.2004090729 Jung, G., Hammer III, J.A., 1994. The actin binding site in the tail domain of Dictyostelium myosin IC ( tail homology region 2 ). FEBS Lett. 342, 197–202.
45 Chapter 1 Introduction
Kalwa, H., Storch, U., Demleitner, J., Fiedler, S., Mayer, T., Kannler, M., Fahlbusch, M., Barth, H., Smrcka, A., Hildebrandt, F., Gudermann, T., Dietrich, A., 2015. Phospholipase C Epsilon (PLCε) Induced TRPC6 Activation: A Common but Redundant Mechanism in Primary Podocytes. J. Cell. Physiol. 230, 1389–1399. doi:10.1002/jcp.24883 Kambara, T., Ikebe, M., 2006. A unique ATP hydrolysis mechanism of single-headed processive myosin, myosin IX. J. Biol. Chem. 281, 4949–4957. doi:10.1074/jbc.M509141200 Kanwar, Y.S., Danesh, F.R., Chugh, S.S., 2007. Contribution of proteoglycans towards the integrated functions of renal glomerular capillaries: a historical perspective. Am. J. Pathol. 171, 9–13. doi:10.2353/ajpath.2007.070356 Kao, W.H.L., Klag, M.J., Meoni, L. a, Reich, D., Berthier-Schaad, Y., Li, M., Coresh, J., Patterson, N., Tandon, A., Powe, N.R., Fink, N.E., Sadler, J.H., Weir, M.R., Abboud, H.E., Adler, S.G., Divers, J., Iyengar, S.K., Freedman, B.I., Kimmel, P.L., Knowler, W.C., Kohn, O.F., Kramp, K., Leehey, D.J., Nicholas, S.B., Pahl, M. V, Schelling, J.R., Sedor, J.R., Thornley-Brown, D., Winkler, C. a, Smith, M.W., Parekh, R.S., 2008. MYH9 is associated with nondiabetic end-stage renal disease in African Americans. Nat. Genet. 40, 1185–1192. doi:10.1038/ng.232 Kaplan, J.M., Kim, S.H., North, K.N., Rennke, H., Correia, L. a, Tong, H.Q., Mathis, B.J., Rodríguez-Pérez, J.C., Allen, P.G., Beggs, a H., Pollak, M.R., 2000. Mutations in ACTN4, encoding alpha-actinin-4, cause familial focal segmental glomerulosclerosis. Nat. Genet. 24, 251– 256. doi:10.1038/73456 Karle, S.M., Uetz, B., Ronner, V., Glaeser, L., Hildebrandt, F., Fuchshuber, A., 2002. Novel mutations in NPHS2 detected in both familial and sporadic steroid-resistant nephrotic syndrome. J. Am. Soc. Nephrol. 13, 388–393. Kestilä, M., Lenkkeri, U., Männikkö, M., Lamerdin, J., McCready, P., Putaala, H., Ruotsalainen, V., Morita, T., Nissinen, M., Herva, R., Kashtan, C.E., Peltonen, L., Holmberg, C., Olsen, a, Tryggvason, K., 1998. Positionally cloned gene for a novel glomerular protein-- nephrin--is mutated in congenital nephrotic syndrome. Mol. Cell 1, 575–82. Kim, J.M., Wu, H., Green, G., Winkler, C. a, Kopp, J.B., Miner, J.H., Unanue, E.R., Shaw, A.S., 2003. CD2-associated protein haploinsufficiency is linked to glomerular disease susceptibility. Science 300, 1298–1300. doi:10.1126/science.1081068 Kim, S.C., Kellett, T., Wang, S., Nishi, M., Nagre, N., Zhou, B., Flodby, P., Shilo, K., Ghadiali, S.N., Takeshima, H., Hubmayr, R.D., Zhao, X., 2014. TRIM72 is required for effective repair of alveolar epithelial cell wounding. AJP Lung Cell. Mol. Physiol. 307, L449–L459. doi:10.1152/ajplung.00172.2014 Kim, S. V, Mehal, W.Z., Dong, X., Heinrich, V., Pypaert, M., Mellman, I., Dembo, M., Mooseker, M.S., Wu, D., Flavell, R. a, 2006. Modulation of cell adhesion and motility in the immune system by Myo1f. Science 314, 136–9. doi:10.1126/science.1131920 Kopp, J.B., Smith, M.W., Nelson, G.W., Johnson, R.C., Freedman, B.I., Bowden, D.W., Oleksyk, T., McKenzie, L.M., Kajiyama, H., Ahuja, T.S., Berns, J.S., Briggs, W., Cho, M.E., Dart, R. a, Kimmel, P.L., Korbet, S.M., Michel, D.M., Mokrzycki, M.H., Schelling, J.R., Simon, E., Trachtman, H., Vlahov, D., Winkler, C. a, 2008. MYH9 is a major-effect risk gene for focal segmental glomerulosclerosis. Nat. Genet. 40, 1175–1184. doi:10.1038/ng.226 Kos, C.H., Le, T.C., Sinha, S., Henderson, J.M., Kim, S.H., Sugimoto, H., Kalluri, R., Gerszten, R.E., Pollak, M.R., 2003. Mice deficient in alpha-actinin-4 have severe glomerular disease. J. Clin. Invest. 111, 1683–1690. doi:10.1172/JCI17988 Kreidberg, J. a, Sariola, H., Loring, J.M., Maeda, M., Pelletier, J., Housman, D., Jaenisch, R., 1993. WT-1 is required for early kidney development. Cell 74, 679–691. doi:10.1016/0092-8674(93)90515-R Krendel, M., Kim, S. V, Willinger, T., Wang, T., Kashgarian, M., Flavell, R. a, Mooseker, M.S., 2009. Disruption of Myosin 1e promotes podocyte injury. J. Am. Soc. Nephrol. 20, 86–94. doi:10.1681/ASN.2007111172 Krendel, M., Mooseker, M.S., 2005. Myosins: tails (and heads) of functional diversity. Physiology (Bethesda). 20, 239–51. doi:10.1152/physiol.00014.2005
46 Chapter 1 Introduction
Krendel, M., Osterweil, E.K., Mooseker, M.S., 2007. Myosin 1E interacts with synaptojanin-1 and dynamin and is involved in endocytosis. FEBS Lett. 581, 644–50. doi:10.1016/j.febslet.2007.01.021 Kriz, W., 2007. Ontogenetic development of the filtration barrier. Nephron. Exp. Nephrol. 106, e44–50. doi:10.1159/000101792 Kuure, S., Vuolteenaho, R., Vainio, S., 2000. Kidney mrophrogenesis: cellular and molecular regulation. Mech Dev 92, 31–45. Lee, H.K., Han, K.H., Jung, Y.H., Kang, H.G., Moon, K.C., Ha, I.S., Choi, Y., Cheong, H. Il, 2011. Variable renal phenotype in a family with an INF2 mutation. Pediatr. Nephrol. 26, 73– 76. doi:10.1007/s00467-010-1644-5 Li, C., Ruotsalainen, V., Tryggvason, K., Shaw, a S., Miner, J.H., 2000. CD2AP is expressed with nephrin in developing podocytes and is found widely in mature kidney and elsewhere. Am. J. Physiol. Renal Physiol. 279, F785–F792. Liao, W., Elfrink, K., Bähler, M., 2010. Head of myosin IX binds calmodulin and moves processively toward the plus-end of actin filaments. J. Biol. Chem. 285, 24933–24942. doi:10.1074/jbc.M110.101105 Lin, P., Zhu, H., Cai, C., Wang, X., Cao, C., Xiao, R., Pan, Z., Weisleder, N., Takeshima, H., Ma, J., 2012. Nonmuscle myosin IIA facilitates vesicle trafficking for MG53- mediated cell membrane repair. FASEB J. 1–9. doi:10.1096/fj.11-188599 Liu, L., Doné, S.C., Khoshnoodi, J., Bertorello, a, Wartiovaara, J., Berggren, P.O., Tryggvason, K., 2001. Defective nephrin trafficking caused by missense mutations in the NPHS1 gene: insight into the mechanisms of congenital nephrotic syndrome. Hum. Mol. Genet. 10, 2637– 2644. Löfgren, M., Ekblad, E., Morano, I., Arner, A., 2003. Nonmuscle Myosin motor of smooth muscle. J. Gen. Physiol. 121, 301–310. doi:10.1085/jgp.200208720 Löwik, M.M., Groenen, P.J.T. a, Pronk, I., Lilien, M.R., Goldschmeding, R., Dijkman, H.B., Levtchenko, E.N., Monnens, L. a, van den Heuvel, L.P., 2007. Focal segmental glomerulosclerosis in a patient homozygous for a CD2AP mutation. Kidney Int. 72, 1198–1203. doi:10.1038/sj.ki.5002469 Machuca, E., Benoit, G., Antignac, C., 2009. Genetics of nephrotic syndrome: Connecting molecular genetics to podocyte physiology. Hum. Mol. Genet. 18, 185–194. doi:10.1093/hmg/ddp328 Männikkö, M., Kestailä, M., Holmberg, C., Norio, R., Ryynänen, M., Olsen, a, Peltonen, L., Tryggvason, K., 1995. Fine mapping and haplotype analysis of the locus for congenital nephrotic syndrome on chromosome 19q13.1. Am. J. Hum. Genet. 57, 1377–1383. Mao, J., Zhang, Y., Du, L., Dai, Y., Gu, W., Liu, A., Shang, S., Liang, L., 2007. NPHS1 and NPHS2 gene mutations in chinese children with sporadic nephrotic syndrome. Pediatr. Res. 61, 117–122. doi:10.1203/01.pdr.0000250041.19306.3d McKenzie, L.M., Hendrickson, S.L., Briggs, W. a, Dart, R. a, Korbet, S.M., Mokrzycki, M.H., Kimmel, P.L., Ahuja, T.S., Berns, J.S., Simon, E.E., Smith, M.C., Trachtman, H., Michel, D.M., Schelling, J.R., Cho, M., Zhou, Y.C., Binns-Roemer, E., Kirk, G.D., Kopp, J.B., Winkler, C. a, 2007. NPHS2 variation in sporadic focal segmental glomerulosclerosis. J. Am. Soc. Nephrol. 18, 2987–2995. doi:10.1681/ASN.2007030319 Melchionda, S., Ahituv, N., Bisceglia, L., Sobe, T., Glaser, F., Rabionet, R., Arbones, M.L., Notarangelo, a, Di Iorio, E., Carella, M., Zelante, L., Estivill, X., Avraham, K.B., Gasparini, P., 2001. MYO6, the human homologue of the gene responsible for deafness in Snell’s waltzer mice, is mutated in autosomal dominant nonsyndromic hearing loss. Am. J. Hum. Genet. 69, 635–640. doi:10.1086/323156 Mele, C., D, B.S., Iatropoulos, P., Donadelli, R., Calabria, A., Eng, D., Maranta, R., Cassis, P., Ph, D., Buelli, S., Tomasoni, S., Piras, R., D, C.P., Krendel, M., Bettoni, S., Biotech, D., Morigi, M., Delledonne, M., Pecoraro, C., Abbate, I., Capobianchi, M.R., Hildebrandt, F., Otto, E., Schaefer, F., Macciardi, F., Ozaltin, F., Emre, S., Ibsirlioglu, T.,
47 Chapter 1 Introduction
Benigni, A., Remuzzi, G., 2011. MYO1E mutations and childhood familial focal segmental glomerulosclerosis. N. Engl. J. Med. 365, 295–306. Miner, J.H., 2012. The glomerular basement membrane. Exp. Cell Res. 318, 973–978. doi:10.1016/j.yexcr.2012.02.031.The Miner, J.H., 1999. Renal basement membrane components. Kidney Int 56, 2016–2024. doi:10.1046/j.1523-1755.1999.00785.x Mollet, G., Ratelade, J., Boyer, O., Muda, A.O., Morisset, L., Lavin, T.A., Kitzis, D., Dallman, M.J., Bugeon, L., Hubner, N., Gubler, M.-C., Antignac, C., Esquivel, E.L., 2009. Podocin inactivation in mature kidneys causes focal segmental glomerulosclerosis and nephrotic syndrome. J. Am. Soc. Nephrol. 20, 2181–2189. doi:10.1681/ASN.2009040379 Morano, I., Chai, G.X., Baltas, L.G., Lamounier-Zepter, V., Lutsch, G., Kott, M., Haase, H., Bader, M., 2000. Smooth-muscle contraction without smooth-muscle myosin. Nat. Cell Biol. 2, 371–375. doi:10.1038/35014065 Morris, S.M., Arden, S.D., Roberts, R.C., Kendrick-Jones, J., Cooper, J. a, Luzio, J.P., Buss, F., 2002. Myosin VI binds to and localises with Dab2, potentially linking receptor- mediated endocytosis and the actin cytoskeleton. Traffic 3, 331–341. doi:tra030503 [pii] Morrison, A. a, Viney, R.L., Saleem, M. a, Ladomery, M.R., 2008. New insights into the function of the Wilms tumor suppressor gene WT1 in podocytes. Am. J. Physiol. Renal Physiol. 295, F12–F17. doi:10.1152/ajprenal.00597.2007 Mount, D.B., 2014. Thick Ascending Limb of the Loop of Henle. Clin. J. Am. Soc. Nephrol. 9, 1974–1986. doi:10.2215/CJN.04480413 Müller, T., Rumpel, E., Hradetzky, S., Bollig, F., Wegner, H., Blumenthal, A., Greinacher, A., Endlich, K., Endlich, N., 2011. Non-muscle myosin IIA is required for the development of the zebrafish glomerulus. Kidney Int. 80, 1055–63. doi:10.1038/ki.2011.256 Muto, S., Hata, M., Taniguchi, J., Tsuruoka, S., Moriwaki, K., Saitou, M., Furuse, K., Sasaki, H., Fujimura, A., Imai, M., Kusano, E., Tsukita, S., Furuse, M., 2010. Claudin-2- deficient mice are defective in the leaky and cation-selective paracellular permeability properties of renal proximal tubules. Proc. Natl. Acad. Sci. U. S. A. 107, 8011–8016. doi:10.1073/pnas.0912901107 Nambiar, R., McConnell, R.E., Tyska, M.J., 2009. Control of cell membrane tension by myosin-I. Proc. Natl. Acad. Sci. U. S. A. 106, 11972–11977. doi:10.1073/pnas.0901641106 Nishikawa, M., Nishikawa, S., Inoue, A., Iwane, A.H., Yanagida, T., Ikebe, M., 2006. A unique mechanism for the processive movement of single-headed myosin-IX. Biochem. Biophys. Res. Commun. 343, 1159–1164. doi:10.1016/j.bbrc.2006.03.057 Nishikawa, S., Homma, K., Komori, Y., Iwaki, M., Wazawa, T., Hikikoshi Iwane, A., Saito, J., Ikebe, R., Katayama, E., Yanagida, T., Ikebe, M., 2002. Class VI myosin moves processively along actin filaments backward with large steps. Biochem. Biophys. Res. Commun. 290, 311–317. doi:10.1006/bbrc.2001.6142 Ohse, T., Chang, A.M., Pippin, J.W., Jarad, G., Hudkins, K.L., Alpers, C.E., Miner, J.H., Shankland, S.J., 2009. A new function for parietal epithelial cells: a second glomerular barrier. Am. J. Physiol. Renal Physiol. 297, F1566–F1574. doi:10.1152/ajprenal.00214.2009 Olsen, A. S., Georgescu, A, Johnson, S., Carrano, A. V, 1996. Assembly of a 1-Mb restriction-mapped cosmid contig spanning the candidate region for Finnish congenital nephrosis (NPHS1) in 19q13.1. Genomics 34, 223–225. doi:10.1006/geno.1996.0270 Ouderkirk, J.L., Krendel, M., 2014. Myosin 1e is a component of the invadosome core that contributes to regulation of invadosome dynamics. Exp. Cell Res. 322, 265–276. doi:10.1016/j.biotechadv.2011.08.021.Secreted Palmer, R.E., Kotsianti, A., Cadman, B., Boyd, T., Gerald, W., Haber, D. a., 2001. WT1 regulates the expression of the major glomerular podocyte membrane protein Podocalyxin. Curr. Biol. 11, 1805–1809. doi:10.1016/S0960-9822(01)00560-7 Patrakka, J., Kestila, M., Wartiovaara, J., Ruotsalainen, V., Tissari, P., Lenkkeri, U., Mannikko, M., Visapaa, I., Holmberg, C., Rapola, J., Tryggvason, K., Jalanko, H., 2000.
48 Chapter 1 Introduction
Congenital nephrotic syndrome (NPHS1): Features resulting from different mutations in Finnish patients. Kidney Int. 58, 972–980. doi:10.1046/j.1523-1755.2000.00254.x Pavenstädt, H., Kriz, W., Kretzler, M., 2003. Cell biology of the glomerular podocyte. Physiol. Rev. 83, 253–307. doi:10.1152/physrev.00020.2002 Philippe, a, Weber, S., Esquivel, E.L., Houbron, C., Hamard, G., Ratelade, J., Kriz, W., Schaefer, F., Gubler, M.-C., Antignac, C., 2008. A missense mutation in podocin leads to early and severe renal disease in mice. Kidney Int. 73, 1038–1047. doi:10.1038/ki.2008.27 Pierchala, B. a, Muñoz, M.R., Tsui, C.C., 2010. Proteomic analysis of the slit diaphragm complex: CLIC5 is a protein critical for podocyte morphology and function. Kidney Int. 78, 868–82. doi:10.1038/ki.2010.212 Pollak, M.R., 2014. Familial FSGS. Adv. Chronic Kidney Dis. 21, 422–425. doi:10.1053/j.ackd.2014.06.001 Putaala, H., Soininen, R., Kilpeläinen, P., Wartiovaara, J., Tryggvason, K., 2001. The murine nephrin gene is specifically expressed in kidney, brain and pancreas: inactivation of the gene leads to massive proteinuria and neonatal death. Hum. Mol. Genet. 10, 1–8. Quaggin, S.E., Kreidberg, J. a, 2008. Development of the renal glomerulus: good neighbors and good fences. Development 135, 609–20. doi:10.1242/dev.001081 Rantanen, M., Palmén, T., Pätäri, A., Ahola, H., Lehtonen, S., Aström, E., Floss, T., Vauti, F., Wurst, W., Ruiz, P., Kerjaschki, D., Holthöfer, H., 2002. Nephrin TRAP mice lack slit diaphragms and show fibrotic glomeruli and cystic tubular lesions. J. Am. Soc. Nephrol. 13, 1586–1594. doi:10.1097/01.ASN.0000016142.29721.22 Rao, M., Balakrishnan, V.S., 2009. The genetic basis of kidney disease risk in African Americans: MYH9 as a new candidate gene. Am. J. Kidney Dis. 53, 579–583. doi:10.1053/j.ajkd.2009.02.005 Ratelade, J., Lavin, T.A., Muda, A.O., Morisset, L., Mollet, G., Boyer, O., Chen, D.S., Henger, A., Kretzler, M., Hubner, N., Théry, C., Gubler, M.-C., Montagutelli, X., Antignac, C., Esquivel, E.L., 2008. Maternal environment interacts with modifier genes to influence progression of nephrotic syndrome. J. Am. Soc. Nephrol. 19, 1491–1499. doi:10.1681/ASN.2007111268 Recuenco, M.C., Ohmori, T., Tanigawa, S., Taguchi, A., Fujimura, S., Conti, M. a., Wei, Q., Kiyonari, H., Abe, T., Adelstein, R.S., Nishinakamura, R., 2014. Nonmuscle Myosin II Regulates the Morphogenesis of Metanephric Mesenchyme-Derived Immature Nephrons. J. Am. Soc. Nephrol. 26, 1081–1091. doi:10.1681/ASN.2014030281 Reiser, J., Kriz, W., Kretzler, M., Mundel, P., 2000. The glomerular slit diaphragm is a modified adherens junction. J. Am. Soc. Nephrol. 11, 1–8. Reiser, J., Polu, K.R., Möller, C.C., Kenlan, P., Altintas, M.M., Wei, C., Faul, C., Herbert, S., Villegas, I., Avila-Casado, C., McGee, M., Sugimoto, H., Brown, D., Kalluri, R., Mundel, P., Smith, P.L., Clapham, D.E., Pollak, M.R., 2005. TRPC6 is a glomerular slit diaphragm-associated channel required for normal renal function. Nat. Genet. 37, 739–744. doi:10.1038/ng1592 Reizes, O., Barylko, B., Li, C., Südhof, T.C., Albanesi, J.P., 1994. Domain structure of a mammalian myosin Iβ. Proc. Natl. Acad. Sci. U. S. A. 91, 6349–6353. doi:10.1073/pnas.91.14.6349 Rhoades, R., Pflanzer, R., 1995. Chapter 23 -- The Kidney, in: Human Physiology. pp. 704–735. Richards, T. a, Cavalier-Smith, T., 2005. Myosin domain evolution and the primary divergence of eukaryotes. Nature 436, 1113–1118. doi:10.1038/nature03949 Rock, R.S., Rice, S.E., Wells, a L., Purcell, T.J., Spudich, J. a, Sweeney, H.L., 2001. Myosin VI is a processive motor with a large step size. Proc. Natl. Acad. Sci. U. S. A. 98, 13655– 13659. doi:10.1073/pnas.191512398 Rood, I.M., Deegens, J.K.J., Wetzels, J.F.M., 2012. Genetic causes of focal segmental glomerulosclerosis: implications for clinical practice. Nephrol. Dial. Transplant 27, 882–90. doi:10.1093/ndt/gfr771
49 Chapter 1 Introduction
Roselli, S., Gribouval, O., Boute, N., Sich, M., Benessy, F., Attié, T., Gubler, M.-C., Antignac, C., 2002. Podocin localizes in the kidney to the slit diaphragm area. Am. J. Pathol. 160, 131–139. doi:10.1016/S0002-9440(10)64357-X Roselli, S., Heidet, L., Sich, M., Kretzler, M., Gubler, M., Henger, A., Antignac, C., Malades, N., 2004. Early Glomerular Filtration Defect and Severe Renal Disease in Podocin- Deficient Mice Early Glomerular Filtration Defect and Severe Renal Disease in Podocin-Deficient Mice Se. Mol. Cell. Biol. 24, 550–560. doi:10.1128/MCB.24.2.550 Ruotsalainen, V., Patrakka, J., Tissari, P., Reponen, P., Hess, M., Kestilä, M., Holmberg, C., Salonen, R., Heikinheimo, M., Wartiovaara, J., Tryggvason, K., Jalanko, H., 2000. Role of nephrin in cell junction formation in human nephrogenesis. Am. J. Pathol. 157, 1905–1916. doi:10.1016/S0002-9440(10)64829-8 Ryffel, G.U., 2003. What can a frog tell us about human kidney development. Nephron. Exp. Nephrol. doi:10.1159/000071282 Šafaríková, M., Reiterová, J., Šafránková, H., Štekrová, J., Zidková, A., Obeidová, L., Kohoutová, M., Tesar, V., 2013. Mutational analysis of ACTN4, encoding a-actinin 4, in patients with focal segmental glomerulosclerosis using HRM method. Folia Biol. (Czech Republic) 59, 110–115. Sanna-Cherchi, S., Burgess, K.E., Nees, S.N., Caridi, G., Weng, P.L., Dagnino, M., Bodria, M., Carrea, A., Allegretta, M. a, Kim, H.R., Perry, B.J., Gigante, M., Clark, L.N., Kisselev, S., Cusi, D., Gesualdo, L., Allegri, L., Scolari, F., D’Agati, V., Shapiro, L.S., Pecoraro, C., Palomero, T., Ghiggeri, G.M., Gharavi, A.G., 2011. Exome sequencing identified MYO1E and NEIL1 as candidate genes for human autosomal recessive steroid-resistant nephrotic syndrome. Kidney Int. 80, 389–96. doi:10.1038/ki.2011.148 Santoro, D., Caccamo, D., Lucisano, S., Buemi, M., Sebekova, K., Teta, D., Nicola, L. De, 2015. Interplay of Vitamin D , Erythropoiesis , and the Renin-Angiotensin System. Biomed Res. Int. Satchell, S.C., Braet, F., 2009. Glomerular endothelial cell fenestrations: an integral component of the glomerular filtration barrier. Am. J. Physiol. Renal Physiol. 296, F947–F956. doi:10.1152/ajprenal.90601.2008 Saxén, L., Sariola, H., 1987. Early organogenesis of the kidney. Pediatr. Nephrol. 1, 385–392. doi:10.1007/BF00849241 Saxén, L., Sariola, H., Lehtonen, E., 1986. Sequential cell and tissue interactions governing organognesis of the kidney. Anat. Embryol. (Berl). 175, 1–6. Schwarz, K., Simons, M., Reiser, J., Saleem, M. a., Faul, C., Kriz, W., Shaw, a. S., Holzman, L.B., Mundel, P., 2001. Podocin, a raft-associated component of the glomerular slit diaphragm, interacts with CD2AP and nephrin. J. Clin. Invest. 108, 1621–1629. doi:10.1172/JCI200112849 Scott, R., and Quaggin, S. 2015. The cell biology of renal filtration. The Journal of Cell Biology. 209(2):199-210. Self, T., Sobe, T., Copeland, N.G., Jenkins, N. a, Avraham, K.B., Steel, K.P., 1999. Role of myosin VI in the differentiation of cochlear hair cells. Dev. Biol. 214, 331–341. doi:10.1006/dbio.1999.9424 Sellers, J.R., 2000. Myosins: a diverse superfamily. Biochim. Biophys. Acta 1496, 3–22. Shankland, S.J., Anders, H.-J., Romagnani, P., 2013. Glomerular parietal epithelial cells in kidney physiology, pathology, and repair. Curr. Opin. Nephrol. Hypertens. 22, 302–9. doi:10.1097/MNH.0b013e32835fefd4 Sherr, E.H., Joyce, M.P., Greene, L. a., 1993. Mammalian myosin Iα, Iβ, and Iγ: New widely expressed genes of the myosin I family. J. Cell Biol. 120, 1405–1416. doi:10.1083/jcb.120.6.1405 Shih, N.Y., Li, J., Cotran, R., Mundel, P., Miner, J.H., Shaw, A S., 2001. CD2AP localizes to the slit diaphragm and binds to nephrin via a novel C-terminal domain. Am. J. Pathol. 159, 2303–2308. doi:10.1016/S0002-9440(10)63080-5
50 Chapter 1 Introduction
Shih, N.Y., Li, J., Karpitskii, V., Nguyen, a, Dustin, M.L., Kanagawa, O., Miner, J.H., Shaw, A S., 1999. Congenital nephrotic syndrome in mice lacking CD2-associated protein. Science 286, 312–315. doi:10.1126/science.286.5438.312 Si, J., Ge, Y., Zhuang, S., Gong, R., 2010. Inhibiting nonmuscle myosin II impedes inflammatory infiltration and ameliorates progressive renal disease. Lab. Invest. 90, 448–458. doi:10.1038/labinvest.2009.142 Simons, M., Wang, M., McBride, O.W., Kawamoto, S., Yamakawa, K., Gdula, D., Adelstein, R.S., Weir, L., 1991. Human nonmuscle myosin heavy chains are encoded by two genes located on different chromosomes. Circ. Res. 69, 530–539. doi:10.1161/01.RES.69.2.530 Souma, T., Suzuki, N., Yamamoto, M., 2015. Renal erythropoietin-producing cells in health and disease. Front. Physiol. 6, 1–10. doi:10.3389/fphys.2015.00167 Stöffler, H.E., Honnert, U., Bauer, C. a, Höfer, D., Schwarz, H., Müller, R.T., Drenckhahn, D., Bähler, M., 1998. Targeting of the myosin-I myr 3 to intercellular adherens type junctions induced by dominant active Cdc42 in HeLa cells. J. Cell Sci. 111 ( Pt 1, 2779–88. Swailes, N.T., Colegrave, M., Knight, P.J., Peckham, M., 2006. Non-muscle myosins 2A and 2B drive changes in cell morphology that occur as myoblasts align and fuse. J. Cell Sci. 119, 3561–3570. doi:10.1242/jcs.03096 Szent-Györgyi, A.G., 2004. The early history of the biochemistry of muscle contraction. J. Gen. Physiol. 123, 631–641. doi:10.1085/jgp.200409091 Thelen, S., Abouhamed, M., Ciarimboli, G., Edemir, B., Bähler, M., 2015. The Rho GAP Myosin IXa is a regulator of kidney tubule function. Am. J. Physiol. - Ren. Physiol. doi:10.1152/ajprenal.00220.2014 Tonna, S.J., Needham, A., Polu, K., Uscinski, A., Appel, G.B., Falk, R.J., Katz, A., Al- Waheeb, S., Kaplan, B.S., Jerums, G., Savige, J., Harmon, J., Zhang, K., Curhan, G.C., Pollak, M.R., 2008. NPHS2 variation in focal and segmental glomerulosclerosis. BMC Nephrol. 9, 13. doi:10.1186/1471-2369-9-13 Trybus, K.M., 1994. Role of myosin light chains. J Muscle Res Cell Motil. Tryggvason, K., 1999. Unraveling the mechanisms of glomerular ultrafiltration: nephrin, a key component of the slit diaphragm. J. Am. Soc. Nephrol. 10, 2440–2445. Tsukaguchi, H., Sudhakar, A., Le, T.C., Nguyen, T., Yao, J., Schwimmer, J. a., Schachter, A.D., Poch, E., Abreu, P.F., Appel, G.B., Pereira, A.B., Kalluri, R., Pollak, M.R., 2002. NPHS2 mutations in late-onset focal segmental glomerulosclerosis: R229Q is a common disease-associated allele. J. Clin. Invest. 110, 1659–1666. doi:10.1172/JCI200216242 Tyska, M.J., Warshaw, D.M., 2002. The Myosin Power Stroke. Cell Motil. Cytoskeleton 15, 1–15. doi:10.1002/cmc.10014 Uetani, N., Bouchard, M., 2009. Plumbing in the embryo: Developmental defects of the urinary tracts. Clin. Genet. 75, 307–317. doi:10.1111/j.1399-0004.2009.01175.x Van Den Berg, R., Hoogenraad, C.C., 2012. Molecular motors in cargo trafficking and synapse assembly. Adv. Exp. Med. Biol. 970, 173–196. doi:10.1007/978-3-7091-0932-8_8 Van den Hoven, M.J., Wijnhoven, T.J., Li, J.-P., Zcharia, E., Dijkman, H.B., Wismans, R.G., Rops, a L., Lensen, J.F., van den Heuvel, L.P., van Kuppevelt, T.H., Vlodavsky, I., Berden, J.H.M., van der Vlag, J., 2008. Reduction of anionic sites in the glomerular basement membrane by heparanase does not lead to proteinuria. Kidney Int. 73, 278–287. doi:10.1038/sj.ki.5002706 Van Duijn, T.J., Anthony, E.C., Hensbergen, P.J., Deelder, A.M., Hordijk, P.L., 2010. Rac1 recruits the adapter protein CMS/CD2AP to cell-cell contacts. J. Biol. Chem. 285, 20137– 20146. doi:10.1074/jbc.M109.099481 Verma, R., Kovari, I., Soofi, A., Nihalani, D., Patrie, K., Holzman, L.B., 2006. Nephrin ectodomain engagement results in Src kinase activation, nephrin phosphorylation, Nck recruitment, and actin polymerization. J. Clin. Invest. 116, 1346–1359. doi:10.1172/JCI27414 Vicente-Manzanares, M., Ma, X., Adelstein, R.S., Horwitz, A.R., 2009. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 10, 778–790. doi:10.1038/nrm2786
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Vize, P., Woolf, A.S., Bard, J., 2003. Introduction : Embryonic Kidneys and Other Nephrogenic Models, in: The Kidney. pp. 1–6. Wagner, M.C., Barylko, B., Albanesi, J.P., 1992. Tissue distribution and subcellular localization of mammalian myosin I. J. Cell Biol. 119, 163–170. doi:10.1083/jcb.119.1.163 Wagner, M.C., Blazer-Yost, B.L., Boyd-White, J., Srirangam, A., Pennington, J., Bennett, S., 2005. Expression of the unconventional myosin Myo1c alters sodium transport in M1 collecting duct cells. Am. J. Physiol. Cell Physiol. 289, C120–C129. doi:10.1152/ajpcell.00569.2003 Wallace, M.A., 1998. Anatomy and physiology of the kidney. AORN. J. 68(5): 800, 803- 16, 819-20; quiz 821-4. Weber, K.L., Sokac, A.M., Berg, J.S., Cheney, R.E., Bement, W.M., 2004. A microtubule-binding myosin required for nuclear anchoring and spindle assembly. Nature 431, 325–329. doi:10.1038/nature02834 Weins, A., Kenlan, P., Herbert, S., Le, T.C., Villegas, I., Kaplan, B.S., Appel, G.B., Pollak, M.R., 2005. Mutational and Biological Analysis of alpha-actinin-4 in focal segmental glomerulosclerosis. J. Am. Soc. Nephrol. 16, 3694–3701. doi:10.1681/ASN.2005070706 Weins, A., Schlondorff, J.S., Nakamura, F., Denker, B.M., Hartwig, J.H., Stossel, T.P., Pollak, M.R., 2007. Disease-associated mutant alpha-actinin-4 reveals a mechanism for regulating its F-actin-binding affinity. Proc. Natl. Acad. Sci. U. S. A. 104, 16080–16085. doi:10.1073/pnas.0702451104 Wells, a L., Lin, a W., Chen, L.Q., Safer, D., Cain, S.M., Hasson, T., Carragher, B.O., Milligan, R. a, Sweeney, H.L., 1999. Myosin VI is an actin-based motor that moves backwards. Nature 401, 505–508. doi:10.1038/46835 Welsch, T., Endlich, N., Gökce, G., Doroshenko, E., Simpson, J.C., Kriz, W., Shaw, A.S., Endlich, K., 2005. Association of CD2AP with dynamic actin on vesicles in podocytes. Am. J. Physiol. Renal Physiol. 289, F1134–F1143. doi:10.1152/ajprenal.00178.2005 Welsh, G.I., Saleem, M. a, 2011. The podocyte cytoskeleton-key to a functioning glomerulus in health and disease. Nat. Rev. Nephrol. 1–8. doi:10.1038/nrneph.2011.151 Winn, M.P., Conlon, P.J., Lynn, K.L., Farrington, M.K., Creazzo, T., Hawkins, A.F., Daskalakis, N., Kwan, S.Y., Ebersviller, S., Burchette, J.L., Pericak-Vance, M. a, Howell, D.N., Vance, J.M., Rosenberg, P.B., 2005. A mutation in the TRPC6 cation channel causes familial focal segmental glomerulosclerosis. Science 308, 1801–1804. doi:10.1126/science.1106215 Wu, L., Pan, L., Wei, Z., Zhang, M., 2011. Structure of MyTH4-FERM domains in myosin VIIa tail bound to cargo. Science 331, 757–760. doi:10.1126/science.1198848 Xie, P., 2010. A model for processive movement of single-headed myosin-IX. Biophys. Chem. 151, 71–80. doi:10.1016/j.bpc.2010.05.007 Yang, L.E., Maunsbach, A.B., Leong, P.K.K., McDonough, A. a, 2005. Redistribution of myosin VI from top to base of proximal tubule microvilli during acute hypertension. J. Am. Soc. Nephrol. 16, 2890–2896. doi:10.1681/ASN.2005040366 Yu, C., Feng, W., Wei, Z., Miyanoiri, Y., Wen, W., Zhao, Y., Zhang, M., 2009. Myosin VI Undergoes Cargo-Mediated Dimerization. Cell 138, 537–548. doi:10.1016/j.cell.2009.05.030 Yu, Z., Ding, J., Huang, J., Yao, Y., Xiao, H., Zhang, J., Liu, J., Yang, J., 2005. Mutations in NPHS2 in sporadic steroid-resistant nephrotic syndrome in Chinese children. Nephrol. Dial. Transplant. 20, 902–908. doi:10.1093/ndt/gfh769 Yuen, S.L., Ogut, O., Brozovich, F. V, 2009. Nonmuscle myosin is regulated during smooth muscle contraction. Am. J. Physiol. Heart Circ. Physiol. 297, H191–H199. doi:10.1152/ajpheart.00132.2009 Zhang, Y., Conti, M.A., Malide, D., Dong, F., Wang, A., Shmist, Y. a., Liu, C., Zerfas, P., Daniels, M.P., Chan, C.C., Kozin, E., Kachar, B., Kelley, M.J., Kopp, J.B., Adelstein, R.S., 2012. Mouse models of MYH9-related disease: Mutations in nonmuscle myosin II-A. Blood 119, 238–250. doi:10.1182/blood-2011-06-358853
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Zhu, T., Ikebe, M., 1994. A novel myosin I from bovine adrenal gland. FEBS Lett. 339, 31–36. doi:10.1016/0014-5793(94)80378-1 Zhu, T., Sata, M., Ikebe, M., 1996. Functional expression of mammalian myosin I beta: analysis of its motor activity. Biochemistry 35, 513–522. doi:10.1021/bi952053c
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Figure 1.1. Anatomical overview of renal filtration.
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Figure 1.1. Anatomical overview of renal filtration. (A) Diagrammatic representation of nephron distribution in the kidney. Glomeruli, the filtration compartments of nephrons, are found within the kidney cortex. (B) Segmental structure of nephrons. The vascularized glomerulus is found at the proximal end and is connected through a series of renal tubules where urinary filtrate composition is refined through reabsorption and secretion. PT, proximal tubule.
LOH, loop of Henle. DT, distal tubule. CD, collecting duct. (C) Cellular organization of the glomeruli. AA, afferent arteriole. BS, Bowman’s space. PT, proximal tubule. DT, distal tubule. EA, efferent arteriole. PEC, parietal epithelial cell. Pod, podocyte. MC, mesangial cell. GEC, glomerular endothelial cell. (Figure and legend are reproduced with permission from (Scott and Quaggin, 2015).
Copyright Journal of Cell Biology).
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Figure 1.2. Schematic cartoon illustrating the glomerular filtration barrier.
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Figure 1.2. Schematic cartoon illustrating the glomerular filtration barrier.
Podocytes are colored in yellow. The cell body and foot process are indicated in the figure. The glomerular basement membrane is colored in light purple. The fenestrated endothelial cells are colored in red. The lumen of the blood vessel is indicated in the picture. The slit diaphragm between two neighboring foot processes is indicated in blue. Small solutes (colored in magenta) are filtered through the fenestrations of the endothelial cells, and glomerular basement membrane, and finally through podocyte slit diaphragms. The magenta arrows indicate the direction of the filtration. (Figure courtesy of Dr. Mira Krendel, with modifications.
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A
B
C
D
Figure 1.3. The development of kidney visceral epithelial cells (podocytes).
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Figure 1.3. The development of kidney visceral epithelial cells (podocytes).
The immature podocytes are a layer of columnar epithelial cells with apical junctional belt (A). As the development continues (B-C), the podocytes undergo morphological change, and transform to mesenchymal-like cells. At this time, the apical junctions move towards the basolateral side, and the primary processes start to form, which subdivide to foot processes. As the glomerulus matures (D), the foot processes interdigitate with the ones from the neighboring podocytes.
The junctions between neighboring foot processes now have moved close proximity to the glomerular basement membrane, and covered by proteincomplex known as slit diaphragm. (Figure is reproduced with modifications with permission from (Kriz, 2007). Copyright S. Karger AG, Basel)
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Figure 1.4. Molecular structure of the slit diaphragm.
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Figure 1.4. Molecular structure of the slit diaphragm. The space between the neighboring foot processes are called filtration slit, which is covered by the protein complex known as slit diaphragm. This protein complex is consist of, but not limited to, transmembrane proteins nephrin, Neph1, and podocin, cytoplasmic adaptor proteins CD2AP and ZO-1, and actin cytoskeleton associated proteins
INF2, Myo1e and α – actinin 4. (Figure is reproduced with permission from
(D’Agati et al., 2011), Copyright Massachusetts Medical Society.)
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Figure 1.5. Schematic domain structures of myosin family of molecular motors discussed in the text.
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Figure 1.5. Schematic domain structures of myosin family of molecular motors discussed in the text. (Figure is redrawn and modified from (Krendel and Mooseker, 2005) with permission.)
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Table 1.1: Epithelial cell tight junctions in the kidney
Epithelial Cell shape in Tight Physiological role References cells light junction2 of tight junction2 microscopy1 components Parietal Squamous Claudin-1 Limit glomerular (Ohse et al., epithelial Occludin filtrate into the 2009) cells ZO-1 urinary space (Shankland et al., 2013) Visceral Arborized Nephrin, Plasma (Fukasawa epithelial (mature Neph1, ultrafiltration et al., 2009; cells podocytes) Podocin, P- Reiser et al., (Podocytes) cadherin, 2000) CD2AP, Catenins ZO-1, JAM- A, Occludin, Cindulin Proximal Cuboidal ZO-1 Leakier (Muto et al., tubule shaped with Occludin epithelium: 2010) epithelial apical Claudin-2, - Paracellular Na+ cells microvilli 10, -11 reabsorption Loop of Cuboidal ZO-1 Leaky epithelium: (Denker and Henle (descending) Thick Thick ascending: Sabath, epithelial Squamous ascending: paracellular Mg2+ , 2011) cells (ascending) Claudin-10, - Mg2+ and NH4+ (Mount, 16, -19 reabsorption 2014) (-3, -4, -8, - Thin descending: 11 in some Paracellular Na+, species) Cl- reabsorption Distal tubule Cuboidal ZO-1 Tight epithelium: (Denker and epithelial shaped Claudin-4, -6, Reabsorb Na+, K+, Sabath, cells without apical -7, -8, -10 and H+ 2011) microvilli Collecting Cuboidal ZO-1 Tighter epithelium: (Denker and tubule/duct Claudin-4, -6, Aquaporin Sabath, epithelial -7, -8, -10, - ENaC, 2011) cells 14 H+/K+-ATPase
1, See (Wallace, 1998).
2, Podocyte junctions are modified adherens junctions, and also specialized tight junctions (Fukasawa et al., 2009; Reiser et al., 2000).
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Table 1.2: Non-muscle myosins in the kidney
Myosin Molecular Monomer Localization Indicated KO mouse type weight or in the kidney function defects (kDa) multimer Myo1c 110-118 Monomer Prominent in Endocytosis; Tubule proximal Exocytosis; defects; tubules and Hearing loss collecting duct Myo1e ~120 Monomer Podocytes Endocytosis; Nephrotic Cell-cell syndrome; adhesion; FSGS Junction maintenance NMMIIA ~230 Dimer or Podocytes; Endocytosis; Only tetramer Proximal Cell-ECM become tubule adhesion severe nephrotic syndrome upon secondary stimulations Myosin ~145 Monomer Proximal Vesicle Tubule VI or dimer tubule apical trafficking; defects microvilli Endocytosis Myosin ~240 Monomer Proximal Endocytosis Tubular VIIb tubule defects epithelium Myosin ~300 Monomer Proximal Endocytosis Tubular IXa or dimer tubule defects epithelium
65 Chapter 2 Myo1e activity in podocyte junction
Chapter 2
Myosin 1e is a component of the glomerular
slit diaphragm complex that regulates actin
reorganization during cell-cell contact
formation in podocytes
Chapter 2 is adapted from the following article published in 2013: Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M., 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am. J. Physiol. Renal Physiol. 305, 532–544.
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2.1 Attributions and Acknowledgments
This chapter is adapted from the publication in 2013 in the journal of
Amercian Journal of Physiology-Renal Physiology. Most of the format has been modified to fit into this thesis, except the references styles. Permission to use as one of my thesis chapters is granted from the journal.
I designed and performed most of the experiments with the technical help from Chase, SE and Pellenz, CD, including glomerular fractionation, immunofluorescence staining, live-cell imaging, calcium switch experiments and pull down assay. Chase, SE helped generated Myo1e-null mice and Pellenz, CD isolated and subcloned Myo1e-null podocytes from Myo1e-null mice. Dr.
Kurihara, H conducted the electron microscopy expements and prepared the figures. Dr. Fanning, AS supplied MDCK cells for domain mapping expement. Dr.
Krendel, M and I prepared the figures and wrote the manuscript, with input from
Dr. Kurihar and Dr. Fanning.
This work was supported by the NIH award 1R01DK083345 to MK and
NIH RO1DK061397 to ASF. Myo1e construct lacking TH2 was provided by J.
Ouderkirk. We are grateful to the members of Krendel lab and to V. Sirotkin, D.
Pruyne, and J. Amack for critical discussion of the manuscript.
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2.2 Abstract
Glomerular visceral epithelial cells, also known as podocytes, are critical both to normal kidney function and the development of kidney disease. Podocyte actin cytoskeleton and their highly specialized cell-cell junctions (also called slit diaphragm complexes) play key roles in controlling glomerular filtration. Myosin
1e (Myo1e) is an actin-based molecular motor that is expressed in renal glomeruli. Disruption of the Myo1e gene in mice and humans promotes podocyte injury and results in the loss of the integrity of the glomerular filtration barrier.
Here we have used biochemical and microscopic approaches to determine whether Myo1e is associated with the slit diaphragm complexes in glomerular podocytes. Myo1e was consistently enriched in the slit diaphragm fraction during subcellular fractionation of renal glomeruli and colocalized with the slit diaphragm markers in mouse kidney. Live cell imaging studies showed that Myo1e was recruited to the newly formed cell-cell junctions in cultured podocytes, where it colocalized with the actin filament cables aligned with the nascent contacts.
Myo1e-null podocytes expressing FSGS-associated Myo1e mutant (A159P) did not efficiently assemble actin cables along new cell-cell junctions. We have mapped domains in Myo1e that were critical for its localization to cell-cell junctions and determined that the SH3 domain of Myo1e tail interacts with ZO-1, a component of the slit diaphragm complex and tight junctions. These findings suggest that Myo1e represents a component of the slit diaphragm complex and may contribute to regulating junctional integrity in kidney podocytes.
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2.3 Introduction
Myosin 1e (Myo1e, previously known as myo1c) is a member of the myosin superfamily of actin-dependent motor proteins that is expressed in many cell types in humans and mice (2, 24, 48). Like other members of the myosin superfamily, Myo1e contains an N-terminal motor domain responsible for actin binding and ATPase activity, a neck domain that includes a light chain-binding IQ motif, and a tail domain that may determine cargo binding properties and intracellular localization. Myo1e tail domain consists of three tail homology (TH) regions: TH1 domain that binds acidic phospholipids (13), proline-rich TH2 domain, and SH3 domain, also known as TH3, which binds to proline-rich proteins (2, 26).
In the kidney, Myo1e expression is particularly high in glomerular visceral epithelial cells, known as podocytes, which play a key role in the selective filtration of proteins in the renal glomerulus (25). Myo1e knockout mice exhibit defects in renal filtration that can be attributed to podocyte dysfunction (6, 25), and mutations in the MYO1E gene are associated with glomerular disease in humans (36, 45). The precise functional role of Myo1e in glomerular podocytes is unknown.
Podocytes are highly specialized epithelial cells that form unusual, arborized, actin-rich projections, known as foot processes. The intertwined foot processes cover the entire surface of glomerular capillaries and are connected with each other via modified cell-cell junctions, called slit diaphragms. Slit
69 Chapter 2 Myo1e activity in podocyte junction diaphragms are unusual junctional complexes that contain tight junction proteins
(ZO-1, occludin, and JAM-A), adherens junction proteins (P-cadherin), and junctional proteins that are specific to podocytes, such as nephrin, Neph-1, and podocin (14). During renal development, slit diaphragms arise from the tight junctions, which translocate from the apical position, characteristic of cuboidal podocyte precursors, to the basolateral position, typical of mature slit diaphragms
(15, 27, 28, 41). The presence of the intact foot processes and slit diaphragms is necessary for normal filtration since mutations in the components of the podocyte actin cytoskeleton or the slit diaphragm proteins are associated with proteinuria
(3, 21-23). Myo1e localization pattern in mouse glomeruli is similar to that of the foot process and slit diaphragm proteins (6, 25). In earlier studies examining
Myo1e expression in cultured cells, endogenous Myo1e (or its rat homolog,
Myr3) was observed in cell-cell junctions (48, 50, 51). Taken together, the filtration defects of Myo1e-null mice and the localization pattern of Myo1e in the kidney and cultured cells suggest that Myo1e may contribute to cell-cell adhesion, and that some of the defects in Myo1e-null podocytes may arise from disruption of cell contacts.
To test this hypothesis in the current study, we used a subcellular fractionation protocol that allows isolation of the detergent-resistant slit diaphragm (SD) fraction from mouse glomeruli (42). In this earlier study (42),
Myo1e was detected in the slit diaphragm fraction by mass spectrometry. In our experiments, the detergent-insoluble SD fraction was enriched in Myo1e,
70 Chapter 2 Myo1e activity in podocyte junction confirming that Myo1e is tightly associated with the SD proteins. We examined the dynamics of Myo1e recruitment to nascent cell-cell contacts in cultured podocytes and the effects of disease-associated Myo1e mutation on its localization to the junctions and on junctional actin assembly, mapped the regions of Myo1e required for junctional localization, and identified tight junction protein ZO-1 as one of the binding partners of Myo1e.
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2.4 Materials and Methods
Antibodies.
Rabbit anti-Myo1e antibody was previously described (25, 48). The following commercial antibodies were used: mouse monoclonal anti-myc (clone
9E10, BioLegend), rabbit anti-GFP and mouse monoclonal anti-ZO-1
(Invitrogen), rabbit anti-podocin (Sigma). Secondary antibodies labeled with
Alexa-488 and -568 were from Invitrogen.
Expression constructs and cloning.
pEGFP-C1 Myo1e and pEGFP-C1 Myo1e tail constructs were previously described (26), as were myc-tagged ZO-1 constructs (11). Truncated Myo1e constructs in pEGFP-C1 vector (Clontech) were produced using In-Fusion HD
PCR cloning (Clontech). Primers used to subclone TH1 domain surround
AA717-920, TH2 domain - AA913-1060, TH3 domain - AA1052-1109. To verify expression level and molecular weight of the full length and truncated Myo1e constructs, GFP-tagged constructs were expressed in HEK-293 cells and analyzed by immunoblotting against GFP (data not shown). EGFP-Myo1e constructs in pShuttle vector for adenoviral expression (Agilent Technologies) were previously described (36). mCherry-Lifeact and mCherry-ZO-1 (human) adenoviral constructs in the pShuttle vector were prepared according to manufacturer’s instructions. GST-tagged proteins were prepared as described
(26).
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Animal studies.
All animal experiments were conducted in accordance with the protocols approved by the SUNY Upstate Committee for Humane Use of Animals (mice) or by the Animal Care and Use Committee of the Juntendo University School of
Medicine (rats for immunoelectron microscopy). Male Wistar rats (4 to 6 weeks old) and neonatal rats (1 to 2 days old) were obtained from Charles River Japan
(Kanagawa, Japan).
Cell culture, transfection and infection.
Conditionally immortalized mouse kidney podocytes lacking Myo1e were obtained from Myo1e knockout mice (25) expressing a transgene encoding a thermosensitive, interferon-inducible variant of the SV-40-T–antigen
(Immortomouse (19)). Wild type podocytes (39) were a generous gift of Dr. Peter
Mundel, Mass General, Boston, MA. Podocyte isolation was performed following procedure from (38). Individual podocyte clones were obtained by dilution cloning and tested by immunoblotting to verify the lack of Myo1e expression as well as the presence of podocyte markers WT-1, synaptopodin, and podocin.
Madin-Darby canine kidney (MDCK) II Tet-Off cells (clone T23; Clontech) and AD-293 (ATCC) were grown in DMEM containing 10% FBS. Immortalized podocytes were cultured as described (38). Podocyte differentiation was induced by a switch from 33°C to 37° and removal of interferon γ from culture medium; cells were differentiated for 2 weeks prior to experiments. MDCK and HEK-293
73 Chapter 2 Myo1e activity in podocyte junction cells were transfected using JetPEI DNA transfection reagent (Polyplus). Mouse podocytes were infected with adenovirus.
Glomerular purification and isolation of slit diaphragm-containing protein fraction.
Glomeruli were purified from mice using a differential sieving technique
(37). Isolation of slit diaphragm proteins was performed as described (42).
Fractionation of glomeruli to isolate slit diaphragm proteins was performed in eight independent experiments with similar results.
GST pulldown assay.
For pulldown from 293 cells, transfected cells were lysed in lysis buffer containing 50mM Tris-HCl (pH 7.5), 150mM NaCl, 1% Triton X-100, 10% glycerol and complete protease inhibitor (Roche) and centrifuged for 10 min at maximum speed in a table-top microcentrifuge at 4oC. The supernatant was incubated with
GST-tagged proteins and glutathione agarose for 3 hours at 4oC. The bead pellets and unbound proteins were separated by centrifugation. Beads were washed one time with lysis buffer then separated again as above. Finally, both bead pellets and unbound proteins were processed for SDS-PAGE.
Immunoelectron microscopy and immunocytochemistry.
Electron microscopy and immunogold labeling were performed as previously described (29). For immunofluorescence staining, mouse kidney cryosections were prepared and processed as described (7).
Fluorescence microscopy and live cell imaging.
For live-cell imaging, MDCK cells or podocytes were plated onto glass-
74 Chapter 2 Myo1e activity in podocyte junction bottom Mattek dishes (Mattek) (for podocyte plating, dishes were precoated with collagen IV). 24 hours post transfection/infection, cells were observed using
Perkin-Elmer UltraView VoX Spinning Disc Confocal system mounted on a Nikon
Eclipse Ti microscope and equipped with an environmental chamber to maintain cells at 37°C. To observe contact reassembly in a calcium switch assay, podocytes were pre-incubated for 20 minutes in HBSS without calcium and magnesium containing 2 mM EGTA to remove calcium, then transferred into
RPMI with 10% FBS and normal calcium content for 5-10 minutes at 37oC before being placed on the microscope stage.
Image analysis.
All fluorescence intensity measurements were done using ImageJ software (NIH). Quantitative analysis was performed using KaleidaGraph
(Synergy software). The statistical significance was calculated using unpaired
Student t-test with unequal variance.
75 Chapter 2 Myo1e activity in podocyte junction
2.5 Results
2.5.1 Myo1e is a component of the slit diaphragm in the renal glomeruli.
To investigate the possibility that Myo1e is enriched in the glomerular slit diaphragm we used a slit diaphragm purification protocol developed by Pierchala et al. (42) to isolate slit diaphragm complex from glomerular preparations. To determine whether Myo1e was present in the slit diaphragm complex-containing fraction, we performed immunoblotting against endogenous Myo1e. Myo1e was indeed enriched in the glomerular slit diaphragm-containing fraction, along with the slit diaphragm markers podocin and ZO-1 (Fig. 2.1A). Surprisingly, Myo1e was retained in the detergent-resistant slit diaphragm fraction whereas in cultured cell lysates Myo1e is usually soluble in non-ionic detergents (see, for example,
(26)). Thus, Myo1e appeared to interact with the components of the detergent- resistant slit diaphragm complex. Unlike Myo1e, actin-binding protein synaptopodin was primarily present in the cytoplasmic fraction (Fig. 2.1A), indicating that the slit diaphragm isolation procedure was specific for the slit diaphragm components. To examine the localization of Myo1e at the ultrastructural level, immunogold labeling of rat kidney sections was performed using antibodies against Myo1e. In the adult rat kidney, Myo1e localized to the intracellular side of the plasma membrane of podocyte foot processes where it was concentrated at the base of the slit diaphragm (open arrowheads; slit diaphragm is indicated by the arrows)(Fig. 2.1B). In the neonatal kidney, where slit diaphragm precursors are undergoing a gradual translocation from the apical
76 Chapter 2 Myo1e activity in podocyte junction to the basal position, Myo1e localization was more variable. While Myo1e was still enriched in the plasma membrane in the regions corresponding to the slit diaphragms, some immunogold particles labeling Myo1e were positioned more apically than in the adult kidney while others were present close to the glomerular basement membrane (Fig. 2.1C).
2.5.2 Myo1e colocalizes with the slit diaphragm marker ZO-1 in mature and immature glomeruli.
Immunostaining of cryosections of mouse kidneys showed that Myo1e colocalized with the slit diaphragm component ZO-1 in mature glomeruli (Fig.
2.2A). Myo1e staining did not completely overlap with ZO-1 labeling (arrows in
2A point to the regions stained for Myo1e only), indicating that Myo1e is present in podocyte cell bodies in addition to being enriched in the slit diaphragm region.
Immunostaining of Myo1e in immature glomeruli in cryosections of one week old mouse kidneys showed that Myo1e was concentrated at the basal aspect of developing podocytes, where it colocalized with ZO-1 but not with the apical marker podocalyxin (Fig. 2.2B). Thus, during podocyte maturation Myo1e is enriched at the basal surface of podocytes, in the region where the formation of new slit diaphragms is taking place.
2.5.3 Myo1e is recruited to cell-cell contacts in cultured podocytes.
To elucidate the role of Myo1e during remodeling of cell-cell junctions, we used conditionally immortalized podocytes to examine the localization of Myo1e.
Myo1e localized to the cell-cell junctions in wild type podocytes (Fig. 2.2C). In
77 Chapter 2 Myo1e activity in podocyte junction order to determine whether the loss of Myo1e activity resulted in changes in assembly of cell-cell junctions, we expressed either wild type or mutant Myo1e in the Myo1e-null podocytes (obtained from the Myo1e KO mice in our lab).
Myo1e-KO podocytes were infected with adenoviruses encoding GFP-tagged
Myo1e and mCherry-tagged Lifeact, a marker of actin filaments (43). To induce contact remodeling, podocytes were subjected to a calcium switch treatment that induces disassembly of cell-cell junctions by decreasing Ca2+ concentration and reassembly of the junctions by Ca2+ replenishment (16). Cell-cell contacts observed using this model likely represent cell-cell adhesions reminiscent of slit diaphragms since they could be immunostained using anti-ZO-1 antibody (data not shown or see supporting Fig. S2.1). Wild type Myo1e was recruited to the newly formed cell-cell adhesions during recovery from low Ca2+ condition (Fig.
2.3), where it colocalized with the actin filaments labeled by Lifeact. Kymographs in Fig. 2.3B illustrate that Myo1e and actin were recruited to the nascent adhesions at the same time during junction formation.
We also examined localization of a mutant form of Myo1e, Myo1e A159P, during formation of new cell-cell contacts in podocytes (Fig. 2.4). A159P is a missense mutation of a highly conserved amino acid residue in the Myo1e motor domain that is associated with the recessive form of FSGS in humans (36).
When expressed in Myo1e-null podocytes, GFP-Myo1eA159P was slightly enriched in the nascent cell-cell junctions but showed a complete lack of colocalization with the actin filaments (Fig. 2.4B, kymographs).
78 Chapter 2 Myo1e activity in podocyte junction
2.5.4 Cells lacking functional Myo1e exhibit defects in actin assembly along cell- cell contacts and in contact spreading.
Formation of cell-cell contacts in Myo1e-knockout podocytes expressing
GFP-tagged wild type Myo1e was accompanied by the assembly of actin punctae along the cell-cell junction (Fig. 2.5A-C, arrowheads, and Fig. 2.3A). These punctae rapidly fused and elongated to form actin cables along the cell-cell boundary. In addition to the actin filament bundles located along the contact, radially positioned short actin spokes (Fig. 2.5B, white arrows) were also observed. The tips of some of these radial spokes colocalized with the areas of enrichment of Myo1e (Fig. 2.5B, black arrow), and in some cases the spokes elongated as the contacts matured and the cells pulled away from each other, while retaining punctate connections. In contrast to the cells expressing wild type
Myo1e, knockout podocytes expressing Myo1e mutant predominantly formed radial actin spokes rather than longitudinal actin cables (Fig. 2.5D-F, white arrows, and Fig. 2.4A). For each Myo1e construct (wild type and A159P mutant),
8 pairs of cells forming contacts were observed. 7 out of 8 pairs of cells expressing GFP-Myo1e were observed to assemble actin cables along the cell- cell junctions while 5 out of 8 pairs of cells expressing GFP-Myo1eA159P failed to assemble actin along the junction and formed only radial actin spokes. Many of the contacts formed by cells expressing Myo1e mutant were relatively small and failed to spread (Fig. 2.5G), suggesting that the lack of assembly of new actin
79 Chapter 2 Myo1e activity in podocyte junction filament bundles along the junction may correlate with inefficient contact formation.
2.5.5 Myo1e localization to cell-cell junctions requires multiple binding motifs.
To further map the domains of Myo1e that are necessary for its localization to the junctions, we used GFP-tagged Myo1e constructs that lack specific tail domains. Since transfection of podocytes is challenging and production of an adenoviral vector for expression of each truncated construct is very resource-intensive, we used MDCK cells for these domain mapping studies
(Fig. 2.6, and supportive Fig. S2.2). As a quantitative measure of junctional localization, we used the ratio of mean fluorescence intensity of GFP-Myo1e along the cell-cell junction to the mean cytosolic intensity of GFP-Myo1e as an indicator of Myo1e enrichment in the junctions (Fig. 2.6B). Both values were background corrected using mean fluorescence intensity of an adjacent untransfected cell as an indicator of background fluorescence. The junctional fold enrichment score for those constructs that are completely cytosolic would be expected to be approximately 0.5 (the intensity along the junction, which includes both a transfected and an untransfected cell, would be approximately half of the cytosolic intensity in the transfected cell). Indeed, constructs that had diffuse cytoplasmic localization had fold enrichment scores close to 0.5 (Fig. 2.6C).
Full length Myo1e and Myo1e tail alone were localized to the intercellular junctions in MDCK cells (Fig. 2.6A). Myo1e tail showed a stronger enrichment in cell-cell junctions than the full length Myo1e (Fig. 2.6C), suggesting that the
80 Chapter 2 Myo1e activity in podocyte junction motor domain may target a subset of Myo1e molecules to other intracellular structures, reducing its junctional enrichment. Myo1e tail expressed in podocytes also exhibited strong localization to cell-cell junctions (supportive Fig. S2.3), indicating that the behavior of truncated constructs in MDCK cells and podocytes was similar. Deletion of the SH3 domain of Myo1e did not significantly impair
Myo1e localization to the junctions (Fig. 2.6A). However, Myo1e construct lacking the TH2 domain had a completely cytosolic localization. Localization of the TH2 domain alone was also diffuse (Fig. 2.6C), while the construct consisting of TH1 domain along with TH2 domain exhibited junctional localization (Fig.
2.6A), and its enrichment in the junctions was similar to that of the full length
Myo1e (Fig. 2.6C). All Myo1e constructs that were enriched in the junctions also colocalized with actin, as indicated by the line scans of GFP-Myo1e and mCherry-Lifeact intensity (Fig. 2.6A, right panels). As shown in Fig. 2.6C,
Myo1e-FL (full-length), Myo1e-tail, Myo1e-ΔSH3 as well as Myo1e-TH1TH2 all exhibited significant enrichment at the junctions. However, if we deleted TH2 domain from the full length Myo1e or TH1 domain from Myo1e tail, the fold enrichment score dropped significantly (Fig. 2.6C). Therefore, we have concluded that TH1 and TH2 domains together are necessary and sufficient to determine junctional localization of Myo1e.
2.5.6 Myo1e SH3 domain binds to the C-terminal portion of ZO-1.
Since Myo1e SH3 domain was not necessary for Myo1e localization to the junctions, we hypothesized that while the main role for TH1 and TH2 domains
81 Chapter 2 Myo1e activity in podocyte junction may be to target Myo1e to the junctions, the SH3 domain may serve as an effector domain, binding or activating additional proteins following Myo1e recruitment to cell-cell contacts. To identify effector proteins that may bind to
Myo1e, we analyzed known protein components of the slit diaphragm complex and their protein interaction motifs. Based on the presence of proline-rich motifs, we hypothesized that ZO-1 could be a potential binding partner for the SH3 domain of Myo1e. To test this potential interaction, we performed an in vitro pulldown assay using GST-tagged SH3 domain of Myo1e to precipitate myc- tagged ZO-1 (Fig. 2.7A) from HEK-293 cell lysates. As shown in Fig. 2.7B, purified GST-tagged Myo1e-SH3 precipitated full-length ZO-1. We used GST- tagged SH3WK mutant as a negative control since this mutation of a tryptophan to a lysine residue has been shown to disrupt SH3 domain binding to proline-rich targets (26). Neither GST alone nor GST-tagged SH3 mutant showed specific binding to ZO-1, which indicates that this interaction was specific to SH3 domain.
Sequence analysis of ZO-1 identified several proline-rich regions located at the
C-terminal portion of ZO-1, which could represent potential binding motifs for the
SH3 domain of Myo1e (Fig. 2.7A). Indeed, only the proline-rich domain containing C-terminal portion of ZO-1, but not the N-terminal portion of ZO-1, bound to Myo1e (Fig. 2.7B). When fluorescently tagged Myo1e and ZO-1 were expressed in cultured podocytes, they colocalized in cell-cell contacts, further confirming the interaction (Fig. 2.7C).
82 Chapter 2 Myo1e activity in podocyte junction
2.6 Discussion
Previous studies have identified a role for Myo1e in the maintenance of glomerular filtration. Mice lacking Myo1e in podocytes exhibit proteinuria, while patients with homozygous mutations in Myo1e are characterized by FSGS and proteinuria (7, 25, 36). The importance of Myo1e for normal glomerular filtration, the localization pattern of Myo1e in glomeruli, and a prior proteomic study that detected Myo1e in the slit diaphragm fraction (42) led us to investigate a potential association between Myo1e and the slit diaphragm complexes.
Using subcellular fractionation of glomerular lysates, we determined that
Myo1e was associated with the slit diaphragm complexes. Immuno-electron microscopy of renal glomeruli indicated that Myo1e was located along the plasma membrane in the foot processes of podocytes, in regions adjacent to the slit diaphragms. These findings suggest that Myo1e may interact with the protein components of the slit diaphragm complex.
Myo1e was also found to colocalize with the slit diaphragm marker ZO-1 in cryosections of mouse kidneys. While Myo1e was enriched in the regions corresponding to the slit diaphragms, a significant amount of Myo1e staining was detected in the cell bodies of podocytes as well as in other regions distinct from the slit diaphragms or cell-cell junctions (Figures 2, 3). Thus, in addition to being highly concentrated in the slit diaphragm region, Myo1e may be associated with other intracellular structures in podocytes, such as endocytic vesicles and cell- substrate adhesions (25, 49). Intriguingly, during glomerular maturation Myo1e
83 Chapter 2 Myo1e activity in podocyte junction was highly enriched in the basal region of developing podocytes, overlapping with ZO-1, which marks developing slit diaphragm complexes. This observation suggested to us that Myo1e may have an important function during formation or remodeling of cell-cell junctions between podocytes.
Live cell imaging and immunostaining of podocytes in culture revealed the presence of Myo1e in cell-cell contacts; this is consistent with the previous observations of actin-binding proteins localizing to cell-cell junctions in podocytes
(1, 15, 46). During contact reassembly, GFP-tagged Myo1e was recruited to the newly formed cell-cell junctions, where it colocalized with the actin filaments assembling along the cell-cell boundary. The precise molecular composition of these newly formed cell adhesions in podocytes is unknown but immunostaining of cells during recovery from Ca2+ removal indicated that ZO-1 was present at these nascent junctions (supportive Fig 1). We hypothesize that these contacts are similar to the slit diaphragms, although they may lack some of the slit diaphragm proteins that are expressed in intact glomeruli but downregulated in cultured podocytes (8). The contacts formed during the initial recovery from calcium removal may also be similar to the nascent adherens junctions (also called punctum adherens), which contain ZO-1 (54).
Myo1e colocalization with the newly assembled actin structures in cell-cell junctions was disrupted in the presence of the disease-associated motor domain mutation (A159P). Since myosin motor domain is responsible for myosin-actin interactions, the lack of colocalization with the actin punctae during contact
84 Chapter 2 Myo1e activity in podocyte junction assembly may reflect the loss of actin binding in the A159P mutant. Alternatively, the abnormal localization of the A159P mutant of Myo1e may be caused by the misfolding of the mutant protein.
Myo1e-knockout podocytes infected with wild type Myo1e assembled two types of actin structures at the nascent junctions: radial actin spokes and circumferential actin filament cables along the cell-cell boundary. We have not been able to directly observe the transitions between the two types of actin filament structures and, therefore, cannot postulate whether the presence of these structures corresponds to distinct stages of contact assembly. However, a similar transition from radial to circumferential actin organization (along with the transition from vinculin-enriched, radially organized tight and adherens junctions to linear junctions depleted of vinculin) has been observed in a calcium switch assay in MDCK cells (52). Myo1e was preferentially localized along the cell-cell junctions, so that the sites of Myo1e enrichment appeared to provide the framework for localizing and/or stabilizing circumferential actin cables. Cells expressing the mutant form of Myo1e did not assemble circumferential actin cables along the cell-cell boundary as efficiently as the cells expressing wild type
Myo1e. Thus, live cell observations suggest that Myo1e may be involved in the formation or remodeling of cell-cell contacts, including transitions from actin punctae or spokes to circumferential actin cables.
Further mapping of the domains determining Myo1e localization to the junctions was performed using MDCK cells. The SH3 domain of Myo1e was
85 Chapter 2 Myo1e activity in podocyte junction dispensable for Myo1e localization to the cell-cell junctions. On the other hand,
TH1 and TH2 domains of Myo1e were necessary for junctional localization. The motor domain of Myo1e was not required for localization of Myo1e to cell-cell junctions since Myo1e tail was able to localize to cell-cell contacts in both MDCK cells and podocytes. Moreover, we have observed that the tail domain was more highly enriched in the cell-cell contacts than the full length Myo1e (Fig. 2.7C).
This observation is consistent with a previous study showing that the tail of
Dictyostelium myosin IB (a long-tailed class I myosin, similar to Myo1e in terms of tail domain organization) is highly enriched at the plasma membrane (3.6-fold enrichment) vs. the full length myosin IB being moderately enriched at the plasma membrane (1.7-fold enrichment) (4). The difference in localization between the full length and tail only constructs of myosin IB in Dictyostelium appears to be due to the interaction of the full length myosin IB with actin filaments in the cytoplasm, which increases the cytoplasmic (non-membrane) fraction of myosin (4).
This observation is in contrast to the observed defects in the localization of the Myo1eA159P mutant during live cell imaging of podocytes. Thus, the A159P mutation may cause a more severe defect in Myo1e localization compared to the deletion of the motor domain, possibly due to the misfolding of the mutant protein. Therefore, the A159P mutant of Myo1e may be unable to interact with either the junctional complexes or the actin filaments. On the other hand, in the
86 Chapter 2 Myo1e activity in podocyte junction absence of the motor domain, Myo1e tail retains its ability to interact with the junctional proteins and the plasma membrane.
How could the various functional domains of Myo1e contribute to its localization to cell-cell junctions? Since the deletion of the proline-rich TH2 domain was sufficient to abolish myosin localization to cell junctions, this region may interact with Myo1e binding partners within the junctional protein complexes; these partners may include SH3 domain containing proteins that remain to be identified. Alternatively, TH2 domain may include an ATP-insensitive actin- binding site, similar to those identified in the TH2 domains of amoeboid class I myosins (20, 44), which may promote interactions with the actin filaments in junctional complexes. TH1 domain of Myo1e interacts with anionic phospholipids with high affinity, with some preference for PtdIns(4,5)P2 and PtdIns(3,4,5)P3
(13). Both PtdIns(4,5)P2 and PtdIns(3,4,5)P3 are enriched at the plasma membrane of MDCK cells where they play important roles in establishment of epithelial cell polarity and epithelial morphogenesis (33). Hydrolysis of
PtdIns(4,5)P2 by bacterial phosphatase SigD from Salmonella results in the loss of junctional integrity, redistribution of ZO-1, and reorganization of junctional actin filaments in intestinal epithelial cells (34), indicating that phosphoinositides play key roles in regulation of epithelial junctional stability. Thus, TH1 domain binding to specific plasma membrane phospholipids together with TH2 domain interactions with proline-rich motif binding proteins or actin filaments may lead to
87 Chapter 2 Myo1e activity in podocyte junction the enrichment of Myo1e in cell-cell junctions in the presence of both lipid- and protein-based signals for the slit diaphragm assembly.
Finally, we sought to identify junctional proteins that interact with Myo1e
SH3 domain. ZO-1, a known component of the slit diaphragm complex, interacted with Myo1e SH3 domain in a pulldown assay. This interaction was mapped to the proline-rich C-terminal portion of ZO-1. We hypothesize that
Myo1e may help recruit ZO-1 to cell-cell junctions via SH3 domain - proline-rich motif interactions. Previous studies have implicated ZO-1 in regulation of actin organization in cell-cell junctions in epithelial cells (12). Thus, ZO-1 and Myo1e may act together to promote coordinated assembly of junctional complexes and the actin-based structural elements that support them.
We hypothesize that the TH1 and TH2 domains together provide signals for targeting Myo1e to the junctions, while Myo1e motor and SH3 domains may serve as effector domains, recruiting additional proteins, such as actin and ZO-1, to nascent contacts (Fig. 2.8).
This study identified Myo1e, a class I myosin, as a component of cell-cell junctions. Other members of the myosin superfamily have been implicated in regulation of cell-cell contact organization (30). Non-muscle myosin II, which is associated with the contractile actin filament bundles that support adherens junctions, regulates junction assembly, organization, and stability (9, 18, 47).
Other myosins that localize to cell-cell contacts in epithelial cells and regulate junction assembly include members of myosin classes VI, VII, IX, and X (5, 10,
88 Chapter 2 Myo1e activity in podocyte junction
17, 31, 32, 40, 53). Unlike other myosins associated with the cell-cell junctions, class I myosins are unlikely to be involved in the long-range transport of vesicles or proteins along actin filaments due to their lack of processivity. Instead, class I myosins, such as Myo1e, may function as dynamic linkers between the plasma membrane lipids, protein components of cell-cell junctions, and actin filaments
(35). Intriguingly, another class I myosin, myo1c, has been implicated in the targeting of the podocyte junctional protein Neph1 to the plasma membrane (1), highlighting the importance of class I myosins in podocyte functions. The ability of class I myosins to interact with both lipid and protein binding partners places these motor proteins in the key position to be able to regulate renal filtration of proteins via modulation of specialized cell-cell contacts between podocytes.
89 Chapter 2 Myo1e activity in podocyte junction
2.7 Reference
1. Arif E, Wagner MC, Johnstone DB, Wong HN, George B, Pruthi PA, Lazzara MJ, and Nihalani D. Motor protein Myo1c is a podocyte protein that facilitates the transport of slit diaphragm protein Neph1 to the podocyte membrane. Mol Cell Biol 31: 2134-2150, 2011. 2. Bement WM, Wirth JA, and Mooseker MS. Cloning and mRNA expression of human unconventional myosin-IC. A homologue of amoeboid myosins-I with a single IQ motif and an SH3 domain. J Mol Biol 243: 356-363, 1994. 3. Boute N, Gribouval O, Roselli S, Benessy F, Lee H, Fuchshuber A, Dahan K, Gubler MC, Niaudet P, and Antignac C. NPHS2, encoding the glomerular protein podocin, is mutated in autosomal recessive steroid-resistant nephrotic syndrome. Nat Genet 24: 349-354, 2000. 4. Brzeska H, Guag J, Preston GM, Titus MA, and Korn ED. Molecular basis of dynamic relocalization of Dictyostelium myosin IB. J Biol Chem 287: 14923-14936, 2012. 5. Chandhoke SK, and Mooseker MS. A role for myosin IXb, a motor-RhoGAP chimera, in epithelial wound healing and tight junction regulation. Mol Biol Cell 23: 2468-2480, 2012. 6. Chase SE, Encina CV, Stolzenburg LR, Tatum A, Holzman LB, and Krendel M. Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am J Physiol Renal Physiol. 7. Chase SE, Encina CV, Stolzenburg LR, Tatum AH, Holzman LB, and Krendel M. Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am J Physiol Renal Physiol 303: F1099-1106, 2012. 8. Chittiprol S, Chen P, Petrovic-Djergovic D, Eichler T, and Ransom RF. Marker expression, behaviors, and responses vary in different lines of conditionally immortalized cultured podocytes. Am J Physiol Renal Physiol 301: F660-671, 2011. 9. Conti MA, Even-Ram S, Liu C, Yamada KM, and Adelstein RS. Defects in cell adhesion and the visceral endoderm following ablation of nonmuscle myosin heavy chain II-A in mice. J Biol Chem 279: 41263-41266, 2004. 10. Etournay R, Zwaenepoel I, Perfettini I, Legrain P, Petit C, and El-Amraoui A. Shroom2, a myosin-VIIa- and actin-binding protein, directly interacts with ZO-1 at tight junctions. J Cell Sci 120: 2838-2850, 2007. 11. Fanning AS, Jameson BJ, Jesaitis LA, and Anderson JM. The tight junction protein ZO-1 establishes a link between the transmembrane protein occludin and the actin cytoskeleton. J Biol Chem 273: 29745-29753, 1998. 12. Fanning AS, Van Itallie CM, and Anderson JM. Zonula occludens-1 and -2 regulate apical cell structure and the zonula adherens cytoskeleton in polarized epithelia. Mol Biol Cell 23: 577-590, 2012. 13. Feeser EA, Ignacio CM, Krendel M, and Ostap EM. Myo1e binds anionic phospholipids with high affinity. Biochemistry 49: 9353-9360, 2010. 14. Fukasawa H, Bornheimer S, Kudlicka K, and Farquhar MG. Slit diaphragms contain tight junction proteins. J Am Soc Nephrol 20: 1491-1503, 2009. 15. George B, and Holzman LB. Signaling from the podocyte intercellular junction to the actin cytoskeleton. Semin Nephrol 32: 307-318, 2012. 16. Hunt JL, Pollak MR, and Denker BM. Cultured podocytes establish a size-selective barrier regulated by specific signaling pathways and demonstrate synchronized barrier assembly in a calcium switch model of junction formation. J Am Soc Nephrol 16: 1593-1602, 2005. 17. Hyenne V, Louvet-Vallee S, El-Amraoui A, Petit C, Maro B, and Simmler MC. Vezatin, a protein associated to adherens junctions, is required for mouse blastocyst morphogenesis. Dev Biol 287: 180-191, 2005. 18. Ivanov AI, Bachar M, Babbin BA, Adelstein RS, Nusrat A, and Parkos CA. A unique role for nonmuscle myosin heavy chain IIA in regulation of epithelial apical junctions. PLoS One 2: e658, 2007.
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19. Jat PS, Noble MD, Ataliotis P, Tanaka Y, Yannoutsos N, Larsen L, and Kioussis D. Direct derivation of conditionally immortal cell lines from an H-2Kb-tsA58 transgenic mouse. Proc Natl Acad Sci U S A 88: 5096-5100, 1991. 20. Jung G, and Hammer JA, 3rd. The actin binding site in the tail domain of Dictyostelium myosin IC (myoC) resides within the glycine- and proline-rich sequence (tail homology region 2). FEBS Lett 342: 197-202, 1994. 21. Kaplan JM, Kim SH, North KN, Rennke H, Correia LA, Tong HQ, Mathis BJ, Rodriguez-Perez JC, Allen PG, Beggs AH, and Pollak MR. Mutations in ACTN4, encoding alpha-actinin-4, cause familial focal segmental glomerulosclerosis. Nat Genet 24: 251-256, 2000. 22. Kestila M, Lenkkeri U, Mannikko M, Lamerdin J, McCready P, Putaala H, Ruotsalainen V, Morita T, Nissinen M, Herva R, Kashtan CE, Peltonen L, Holmberg C, Olsen A, and Tryggvason K. Positionally cloned gene for a novel glomerular protein--nephrin--is mutated in congenital nephrotic syndrome. Mol Cell 1: 575-582, 1998. 23. Kim JM, Wu H, Green G, Winkler CA, Kopp JB, Miner JH, Unanue ER, and Shaw AS. CD2-associated protein haploinsufficiency is linked to glomerular disease susceptibility. Science 300: 1298-1300, 2003. 24. Kim SV, Mehal WZ, Dong X, Heinrich V, Pypaert M, Mellman I, Dembo M, Mooseker MS, Wu D, and Flavell RA. Modulation of cell adhesion and motility in the immune system by Myo1f. Science 314: 136-139, 2006. 25. Krendel M, Kim SV, Willinger T, Wang T, Kashgarian M, Flavell RA, and Mooseker MS. Disruption of Myosin 1e promotes podocyte injury. J Am Soc Nephrol 20: 86-94, 2009. 26. Krendel M, Osterweil EK, and Mooseker MS. Myosin 1E interacts with synaptojanin-1 and dynamin and is involved in endocytosis. FEBS Lett 581: 644-650, 2007. 27. Kriz W. Ontogenetic development of the filtration barrier. Nephron Exp Nephrol 106: e44- 50, 2007. 28. Kurihara H, Anderson JM, and Farquhar MG. Diversity among tight junctions in rat kidney: glomerular slit diaphragms and endothelial junctions express only one isoform of the tight junction protein ZO-1. Proc Natl Acad Sci U S A 89: 7075-7079, 1992. 29. Kurihara H, Harita Y, Ichimura K, Hattori S, and Sakai T. SIRP-alpha-CD47 system functions as an intercellular signal in the renal glomerulus. Am J Physiol Renal Physiol 299: F517-527, 2010. 30. Liu KC, and Cheney RE. Myosins in cell junctions. Bioarchitecture 2: 2012. 31. Liu KC, Jacobs DT, Dunn BD, Fanning AS, and Cheney RE. Myosin-X functions in polarized epithelial cells. Mol Biol Cell 23: 1675-1687, 2012. 32. Maddugoda MP, Crampton MS, Shewan AM, and Yap AS. Myosin VI and vinculin cooperate during the morphogenesis of cadherin cell cell contacts in mammalian epithelial cells. J Cell Biol 178: 529-540, 2007. 33. Martin-Belmonte F, Gassama A, Datta A, Yu W, Rescher U, Gerke V, and Mostov K. PTEN-mediated apical segregation of phosphoinositides controls epithelial morphogenesis through Cdc42. Cell 128: 383-397, 2007. 34. Mason D, Mallo GV, Terebiznik MR, Payrastre B, Finlay BB, Brumell JH, Rameh L, and Grinstein S. Alteration of epithelial structure and function associated with PtdIns(4,5)P2 degradation by a bacterial phosphatase. J Gen Physiol 129: 267-283, 2007. 35. McConnell RE, and Tyska MJ. Leveraging the membrane - cytoskeleton interface with myosin-1. Trends Cell Biol 20: 418-426, 2010. 36. Mele C, Iatropoulos P, Donadelli R, Calabria A, Maranta R, Cassis P, Buelli S, Tomasoni S, Piras R, Krendel M, Bettoni S, Morigi M, Delledonne M, Pecoraro C, Abbate I, Capobianchi MR, Hildebrandt F, Otto E, Schaefer F, Macciardi F, Ozaltin F, Emre S, Ibsirlioglu T, Benigni A, Remuzzi G, and Noris M. MYO1E mutations and childhood familial focal segmental glomerulosclerosis. N Engl J Med 365: 295-306, 2011. 37. Mundel P, Gilbert P, and Kriz W. Podocytes in glomerulus of rat kidney express a characteristic 44 KD protein. J Histochem Cytochem 39: 1047-1056, 1991.
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38. Mundel P, Heid HW, Mundel TM, Kruger M, Reiser J, and Kriz W. Synaptopodin: an actin-associated protein in telencephalic dendrites and renal podocytes. J Cell Biol 139: 193-204, 1997. 39. Mundel P, Reiser J, Zuniga Mejia Borja A, Pavenstadt H, Davidson GR, Kriz W, and Zeller R. Rearrangements of the cytoskeleton and cell contacts induce process formation during differentiation of conditionally immortalized mouse podocyte cell lines. Exp Cell Res 236: 248- 258, 1997. 40. Omelchenko T, and Hall A. Myosin-IXA regulates collective epithelial cell migration by targeting RhoGAP activity to cell-cell junctions. Curr Biol 22: 278-288, 2012. 41. Pavenstadt H, Kriz W, and Kretzler M. Cell biology of the glomerular podocyte. Physiol Rev 83: 253-307, 2003. 42. Pierchala BA, Munoz MR, and Tsui CC. Proteomic analysis of the slit diaphragm complex: CLIC5 is a protein critical for podocyte morphology and function. Kidney Int 78: 868- 882, 2010. 43. Riedl J, Crevenna AH, Kessenbrock K, Yu JH, Neukirchen D, Bista M, Bradke F, Jenne D, Holak TA, Werb Z, Sixt M, and Wedlich-Soldner R. Lifeact: a versatile marker to visualize F-actin. Nat Methods 5: 605-607, 2008. 44. Rosenfeld SS, and Rener B. The GPQ-rich segment of Dictyostelium myosin IB contains an actin binding site. Biochemistry 33: 2322-2328, 1994. 45. Sanna-Cherchi S, Burgess KE, Nees SN, Caridi G, Weng PL, Dagnino M, Bodria M, Carrea A, Allegretta MA, Kim HR, Perry BJ, Gigante M, Clark LN, Kisselev S, Cusi D, Gesualdo L, Allegri L, Scolari F, D'Agati V, Shapiro LS, Pecoraro C, Palomero T, Ghiggeri GM, and Gharavi AG. Exome sequencing identified MYO1E and NEIL1 as candidate genes for human autosomal recessive steroid-resistant nephrotic syndrome. Kidney Int 2011. 46. Schiwek D, Endlich N, Holzman L, Holthofer H, Kriz W, and Endlich K. Stable expression of nephrin and localization to cell-cell contacts in novel murine podocyte cell lines. Kidney Int 66: 91-101, 2004. 47. Shewan AM, Maddugoda M, Kraemer A, Stehbens SJ, Verma S, Kovacs EM, and Yap AS. Myosin 2 is a key Rho kinase target necessary for the local concentration of E-cadherin at cell-cell contacts. Mol Biol Cell 16: 4531-4542, 2005. 48. Skowron JF, Bement WM, and Mooseker MS. Human brush border myosin-I and myosin-Ic expression in human intestine and Caco-2BBe cells. Cell Motil Cytoskeleton 41: 308- 324, 1998. 49. Soda K, Balkin DM, Ferguson SM, Paradise S, Milosevic I, Giovedi S, Volpicelli- Daley L, Tian X, Wu Y, Ma H, Son SH, Zheng R, Moeckel G, Cremona O, Holzman LB, De Camilli P, and Ishibe S. Role of dynamin, synaptojanin, and endophilin in podocyte foot processes. J Clin Invest 122: 4401-4411, 2012. 50. Stoffler HE, Honnert U, Bauer CA, Hofer D, Schwarz H, Muller RT, Drenckhahn D, and Bahler M. Targeting of the myosin-I myr 3 to intercellular adherens type junctions induced by dominant active Cdc42 in HeLa cells. J Cell Sci 111 ( Pt 18): 2779-2788, 1998. 51. Stoffler HE, Ruppert C, Reinhard J, and Bahler M. A novel mammalian myosin I from rat with an SH3 domain localizes to Con A-inducible, F-actin-rich structures at cell-cell contacts. J Cell Biol 129: 819-830, 1995. 52. Twiss F, Le Duc Q, Van Der Horst S, Tabdili H, Van Der Krogt G, Wang N, Rehmann H, Huveneers S, Leckband DE, and De Rooij J. Vinculin-dependent Cadherin mechanosensing regulates efficient epithelial barrier formation. Biol Open 1: 1128-1140, 2012. 53. Velichkova M, Guttman J, Warren C, Eng L, Kline K, Vogl AW, and Hasson T. A human homologue of Drosophila kelch associates with myosin-VIIa in specialized adhesion junctions. Cell Motil Cytoskeleton 51: 147-164, 2002. 54. Yonemura S. Cadherin-actin interactions at adherens junctions. Curr Opin Cell Biol 23: 515-522, 2011.
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A
Input CytoplasmLight membranesDR slit diaphragmDS slit diaphragm Input Cytoplasm Light membraneDR slit diaphragmDS slit diaphragm ZO-1 Myo1e Myo1e Podocin Synaptopodin B C
Slit diaphragm ∆ ∆ ∆ ∆ ∆ ∆ Foot process
∆ ∆ ∆
∆ ∆ ∆
Immuno-gold ∆ particle
Figure 2.1. Myo1e is a component of slit diaphragm.
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Figure 2.1. Myo1e is a component of slit diaphragm. A, Western blot analysis of glomerular subfractions. Samples were blotted against endogenous ZO-1,
Myo1e, podocin, and synaptopodin. Left and right panels (from two representative fractionation experiments) indicate that Myo1e, podocin, and ZO-1 are enriched in the detergent-resistant (DR) slit diaphragm fraction while another podocyte marker, synaptopodin, is present in the cytoplasmic fraction. DS, detergent soluble. B, Immuno-electron micrograph of the adult rat glomerulus shows that Myo1e is concentrated at the base of the slit diaphragm (left panel).
The same micrograph with the outlines of podocyte foot processes is shown on the right. Arrows indicate slit diaphragm, while arrowheads point to gold particles that mark the anti-Myo1e antibody. C, Immuno-electron micrograph of the neonatal rat glomerulus displays variable location of Myo1e along the lateral surface of podocyte foot processes. Most immunogold particles marking Myo1e position are located adjacent to the slit diaphragms; however, the distance between Myo1e signal and the glomerular basement membrane is highly variable. Bars, 100 nm.
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95 Chapter 2 Myo1e activity in podocyte junction
Figure 2.2. Myo1e colocalizes with the slit diaphragm marker ZO-1 in mouse glomeruli and in cultured podocytes. A, immunostaining of cryosections of two week old mouse kidney with antibodies against ZO-1 and
Myo1e shows colocalization of Myo1e and ZO-1 (yellow in merge) in mature glomeruli as well as areas stained for Myo1e only (green in merge, indicated by arrows). B, immunostaining of cryosections of one week old mouse kidney against podocalyxin, ZO-1 and Myo1e shows that ZO-1 and Myo1e localize to the basal aspect of podocytes in the immature glomeruli while podocalyxin marks the apical side. C, immunostaining of wild type mouse podocytes reveals localization of Myo1e and ZO-1 to cell-cell junctions. Scale bar, 10 µm.
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Figure 2.3. Myo1e is recruited to the nascent contacts in cultured podocytes.
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Figure 2.3. Myo1e is recruited to the nascent contacts in cultured podocytes. A, Myo1e KO podocytes were infected with adenoviruses encoding
GFP-Myo1e and mCherry-Lifeact for 24 hours and subjected to calcium removal and replenishment to induce disassembly and reassembly of cell contacts. Time- lapse imaging was started 5 minutes after the addition of normal calcium- containing medium. Frames selected from a time-lapse movie show Myo1e and
Lifeact localization at 10, 20 and 30 min time points during imaging. Myo1e and actin are recruited to the newly formed contacts. Bars, 10 µm. B, kymographs show that Myo1e and actin are both recruited to the nascent contacts at the same time.
98 Chapter 2 Myo1e activity in podocyte junction A Myo1e-A159P Lifeact Myo1e-A159P Lifeact 10 min
20 min
30 min
B Myo1e-A159P Lifeact
24 min
10 um
Figure 2.4. Myo1eA159P mutant does not colocalize with actin in the newly formed contacts.
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Figure 2.4. Myo1eA159P mutant does not colocalize with actin in the newly formed contacts. A, Myo1e KO podocytes were infected with adenovirus encoding GFP-Myo1eA159P and mCherry-lifeact 24 hours before calcium switch treatment and live cell imaging. Selected frames from a time-lapse movie show cells at 10, 20 and 30 min time points. Bars, 10 µm. B, kymographs show that
Myo1eA159P does not colocalize with nascent actin bundles.
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A B C
Lifeact
Myo1e
DEF
Lifeact
Myo1e A159P
G Rate of contact spreading, µm/min 4 3 2 1 0 -1
WT A159P Figure 2.5. Podocytes expressing mutant Myo1e fail to assemble actin filament bundles along cell-cell boundaries.
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Figure 2.5. Podocytes expressing mutant Myo1e fail to assemble actin filament bundles along cell-cell boundaries. Contact reassembly following calcium switch treatment was observed in Myo1e KO podocytes expressing
GFP-Myo1e and mCherry-lifeact (A, B, C) or GFP-Myo1eA159P and mCherry- lifeact (D, E, F). A-C, cells expressing wild type Myo1e assembled actin filaments along the cell-cell junction (arrowheads) and perpendicular to the junctions (white arrows). Bundles running perpendicular to the junctions were associated with Myo1e-containing punctae at the cell edge (black arrow). Panels
A and B include enlarged images of the areas indicated by white rectangular outlines. D-F, cells expressing Myo1e mutant assembled primarily radial bundles of actin filaments (white arrows). Bars, 10 µm. G, box-and-whiskers plots of the rate of contact spreading measured using 9-10 minute long time-lapse movies of
Myo1e KO podocytes expressing either GFP-Myo1e (N=5 cells) or GFP-
Myo1eA159P (N=6 cells). Horizontal line within each box indicates the median for each group, boxes corresponds to the 25-75 percentile range, and outliers are shown as open circles.
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A mCherry-Lifeact GFP Distance along the line (μm) Actin Myo1e
Actin Myo1e-FL 0 5 10 15 20
Actin Myo1e-tail 0 5 10 15 20
Actin Myo1e-∆SH3 Fluorescence intensity (A.U.) 0 5 10 15 20
Actin Myo1e-∆TH2 0 5 10 15 20 Actin Myo1e-TH1TH2
0 5 10 15 20 25 B C Fold enrichment of myo1e (J-B)/(C-B) Myo1e-FL GFP Motor IQ SH3TH2TH1 J Myo1e-Tail GFP TH1 SH3TH2 * C B Myo1e-∆SH3 GFP Motor IQ TH2TH1 Myo1e-∆TH2 GFP Motor IQ TH1 SH3 * * * * Myo1e-∆TH2SH3 GFP Motor IQ TH1 Myo1e-TH1 GFP TH1 * * * * Myo1e-TH2 GFP TH2 MI - MI GFP SH3 * * Fold junction enrichment = J B Myo1e-SH3 - GFP TH2TH1 MIC MIB Myo1e-TH1TH2 Myo1e-TH2SH3 GFP SH3TH2 * * 02468
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Figure 2.6. Myo1e localization to the junctions requires multiple domain interactions. A, confocal sections through the apical portion of MDCK cells expressing GFP-tagged Myo1e constructs and mCherry-Lifeact. Full length
Myo1e, Myo1e tail, Myo1e lacking SH3 domain, and Myo1e construct consisting of TH1 and TH2 domains localize to the junctions while Myo1e lacking TH2 domain shows primarily cytosolic localization. Scale bar, 10 µm. Line scans in the right hand panels show fluorescence intensity of the GFP and mCherry signals along a line drawn across two of the cell-cell junctions (indicated by the white lines in merged images). Peaks of GFP and mCherry fluorescence at the junctions coincide, except for the Myo1e construct lacking TH2 domain, which is primarily cytosolic. B, method used to calculate fold enrichment of Myo1e in the junctions. A line drawn along the junction of a transfected cell (green) with an untransfected cell (white) was used to measure mean fluorescence intensity (MI) in the junction (J) while a line drawn in the cytosol of a transfected cell was used to measure mean fluorescence intensity in the cytosol (C). Mean fluorescence intensity of the background (B) was measured by drawing a line through an untransfected cell. C, analysis of the fold junctional enrichment of Myo1e. The bar graphs represent mean fold enrichment for 10 cells per construct (16 cells for the full length Myo1e) while the error bars correspond to standard deviation.
Asterisks indicate constructs that showed statistically significant difference in fold enrichment relative to the full length Myo1e, with P-values less than 0.01. Double
104 Chapter 2 Myo1e activity in podocyte junction asterisks indicate P-values less than 0.0001. P-values for the t-test comparison to the full length Myo1e: DeltaSH3 = 0.2957, TH1TH2 = 0.01135.
105 Chapter 2 Myo1e activity in podocyte junction
A Myo1e-FL Motor IQ TH1 TH2SH3
GST-Myo1e-SH3 GST SH3
GST GST
GST-Myo1e-SH3wk GST SH3wk
ZO-1 FL PDZ1 PDZ2 PDZ3 SH3 GUK U6 α ABR ZU5
ZO-1 N-term PDZ1 PDZ2 PDZ3 SH3 GUK U6 (1-888) ZO-1 C-term ABR ZU5 (∆79-1045)
B Input Unbound Bound
GST-SH3 + + GST + + GST-SH3wk + + Myc-ZO-1 FL
Myc-ZO-1 N-term WB: anti-myc (1-888)
Myc-ZO-1 C-term (Δ79-1045) C
GFP-myo1e mCherry-ZO-1
Figure 2.7. Myo1e SH3 domain interacts with the C terminal portion of ZO-
1.
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Figure 2.7. Myo1e SH3 domain interacts with the C terminal portion of ZO-
1. A, a diagram showing domain organization of Myo1e and ZO-1 constructs used in the pulldown experiments. All ZO-1 constructs were tagged with the myc tag. Protein domains are indicated by the following abbreviations: TH = Tail
Homology; SH3 – Src Homology 3; PDZ = PSD-95/Dlg1/ZO-1; GUK =GUanylate
Kinase; U6 = Unique 6; ZU5 =ZO-1, UNC-5 proline-rich. B, Western blot analysis of GST pulldown reactions using anti-myc antibody. GST-SH3 domain of Myo1e but not GST alone or GST-SH3 mutant precipitated myc-tagged ZO-1 full length and ZO-1 C-terminus. C, GFP-labeled Myo1e and mCherry-tagged
ZO-1 (full length) colocalize when expressed in mouse podocytes. Scale bar, 10
µm.
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A B
Myo1e motor domain TH2 ZO-1 TH1 Actin SH3
Figure 2.8. Model describing how Myo1e may function in cell-cell junctions.
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Figure 2.8. Model describing how Myo1e may function in cell-cell junctions.
A, Myo1e localization to the junctions requires multiple domain interactions.
Interactions of the TH1 domain with the plasma membrane lipids and binding of the proline-rich TH2 domain to junction-associated proteins target Myo1e to the junctions. B, motor domain of Myo1e interacting with actin filaments and SH3 domain interacting with proline-rich proteins, such as ZO-1, promote coordinated assembly of cell-cell junctions and junctional actin bundles.
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Supplementary Figure S2.1. Localization of endogenous Myo1e and ZO-1 after recovery from a calcium switch assay.
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Supplementary Figure S2.1. Localization of endogenous Myo1e and ZO-1 after recovery from a calcium switch assay. Wild type podocytes were placted on coverslips and treated with HBSS for 20 min at 37oC to allow disassembly of the junctions. Fresh medium (RPMI+10%FBS+ABM) then added after removal of
HBSS for 30 min to allow the junctions to reassemble. Coverslips were then fixed and stained against endogenous Myo1e and ZO-1. Scale bars: 10 µm.
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Supplementary Figure S2.2. Localization of Myo1e constructs in MDCK cells.
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Supplementary Figure S2.2. Localization of Myo1e constructs in MDCK cells. (A) Confocal images of MDCK cells transfected with GFP-tagged Myo1e constructs and mCherry-tagged Lifeact. Scale bars, 10 µm. (B) Western blot analysis of different GFP-tagged Myo1e construct from 293 cells lysate.
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Supplementary Figure S2.3. Tail domain of Myo1e localizes to cell-cell contacts in podocytes.
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Supplementary Figure S2.3. Tail domain of Myo1e localizes to cell-cell contacts in podocytes. Confocal sections of Myo1e-KO podocytes infected with adenoviruses encoding GFP-Myo1e-tail and mCherry-Lifeact. Scale bars,
10 µm.
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Chapter 3
Dissecting the effects of kidney disease-
associated Myo1e motor domain mutants on
junction dynamics
Chapter 3 is written by myself and edited by my colleagues for grammatical errors. I designed and performed all the experiments, analyzed all the data, and prepared all the figures. This chapter is not yet ready for publication, since there are still a few more experiments to perform in order to finish the story.
116 Chapter 3 Myo1e motor domain mutants activity in podocytes
3.1 Attributions and Acknowledgments
I want to thank the technicians and graduate student in my lab for their long term technical support and help. I also want to thank my advisor Dr. Mira
Krendel, and all of the Weiskotten Hall basement labs for critical discussion of the project.
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3.2 Abstract
Focal segmental glomerulosclerosis (FSGS) is the most common primary renal disorder that will lead to end-stage kidney disease, such as renal failure, if not treated properly. The pathological features of FSGS include scar tissue formation in both focal and segmental portions of the glomeruli, and massive proteinuria. In the past four decades, increased research has provided evidence that FSGS is the disease of glomerular podocytes, which are being attacked during pathological processes. Podocytes are specialized epithelial cells in renal glomeruli, which cover the surface of the capillaries. They play an essential role in maintaining proper renal filtration function. Among all genetic causes of FSGS, mutations in MYO1E gene, A159P and T119I, were found to be associated with childhood familial FSGS. Myosin 1e (myo1e) is a non-muscle myosin, which has a relatively high expression profile in kidney podocytes in both humans and mice.
It has been found to be a component of the slit diaphragms, specialized junctions between podocyte foot processes, which plays a key role in maintaining slit diaphragm integrity. However, the molecular mechanism that links myosin 1e and
FSGS still remains unclear.
Using immortalized myo1e-null cultured podocytes that isolated from mice kidneys, we were able to mimic cell-cell junction assembly upon calcium-switch experiments. We introduced GFP-tagged myo1e constructs into cultured podocytes using adenovirus-encoded vectors and utilized live-cell imaging to visualize the recruitment of junctional components during junction assembly.
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We found that without functional myo1e, cultured podocytes were unable to assemble proper junctional structures. Furthermore, FSGS-associated myo1e mutants do not recruit to the nascent junctions simultaneously with other junctional components, such as actin and ZO-1. Therefore, we have concluded that mutations in FSGS-associated MYO1E gene will result in defect during cell- cell junction assembly, which mimic the “leaky” slit diaphragm in vivo.
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3.3 Introduction
Focal segmental glomerulosclerosis (FSGS) is a common primary kidney disorder causing end-stage kidney disease in the United States (D’Agati et al.,
2011). The defining feature of FSGS is massive proteinuria (protein loss from the urine) with morphological and pathological changes of the renal filtration barrier
(D’Agati et al., 2011). Kidney visceral epithelial cells, also known as podocytes, are the key component of the glomerular filtration barrier. Podocytes are specialized epithelial cells with unique cellular structures. They form long protrusions called foot processes that interdigitate with the foot processes of neighboring podocytes, and cover blood vessels (Faul et al., 2007). The spaces between adjacent foot processes are called filtration slits, which consist of protein complexes such as transmembrane proteins and cytoplasmic adaptor proteins, and are known as slit diaphragms (D’Agati et al., 2011; Welsh and Saleem,
2011). The actin cytoskeleton within the foot processes plays an essential role to support podocyte shape and regulate their dynamics. Disruption of the podocyte actin cytoskeleton is the cause of many pathological processes (Mathieson,
2011; Welsh and Saleem, 2011), and in the past decade many genetic mutations have been discovered that are associated with FSGS (See review Rood et al.,
2012).
Myosin 1e (Myo1e) is a non-muscle myosin that is broadly expressed in the humans and mice, with the highest expression profile in the spleen and lymph nodes (Kim et al., 2006). In the kidney, Myo1e is enriched in the podocytes.
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Genetic studies and high throughput sequencing have identified two mutations
(A159P and T119I) in the motor domain of Myo1e that are associated with childhood familial FSGS (Al-Hamed et al., 2013; Mele et al., 2011; Sanna-
Cherchi et al., 2011). Both mutations are missense mutations leading to a single amino acid change. In our previous study (see chapter 2), we found that Myo1e is a component of the slit diaphragm and the A159P mutation in the motor domain affects actin reorganization during cell-cell contact formation (Bi et al.,
2013). In this chapter, we are focusing on the other motor domain mutant, T119I, and investigating the effects of T119I mutant on podocytes junction dynamics.
The glomerular slit diaphragm is a modified adherens junction (Reiser et al., 2000), and also contains tight junction proteins (Fukasawa et al., 2009). Our previous study has shown the co-localization of tight junction protein ZO-1 and
Myo1e in the rat kidney glomerulus via immunostaining, and the interactions between the Myo1e SH3 domain and the C-terminal portion of ZO-1 via in vitro pulldown assay (Bi et al., 2013). This makes ZO-1 an ideal marker for the study of the dynamics of the podocytes junctions.
We investigated the junctional dynamics using Myo1e-null immortal podocytes by co-expressing GFP-tagged Myo1e wild type or motor domain mutants, and mCherry-tagged ZO-1 by adenoviral vectors infection. We were able to observe mCherry-tagged ZO-1 recruitment in cells expressing different
Myo1e constructs. Fluorescence recovery after photobleaching (FRAP) experiments on mCherry-tagged ZO-1 at podocytes junctions showed that
121 Chapter 3 Myo1e motor domain mutants activity in podocytes junctional ZO-1 turnover was increased in cells expressing the Myo1e-A159P mutant. This suggests that without functional Myo1e in the cells, the junctions are more dynamic and less stable, revealing a disease model in which loss of Myo1e results in a “leaky” slit diaphragm.
122 Chapter 3 Myo1e motor domain mutants activity in podocytes
3.4 Material and Methods
Cell culture and infection
Conditionally immortalized mouse kidney podocytes lacking Myo1e were obtained from Myo1e knockout mice (Krendel et al., 2009) expressing a transgene encoding a thermosensitive, interferon-inducible variant of the SV-40-T antigen (Immortomouse (Jat et al., 1991)). The isolation of the podocyte clones was done as previously described (Bi et al., 2013).
Immortalized Myo1e-null podocytes were cultured as described (Mundel et al., 1997). Podocyte differentiation was induced by a switch from 33oC to 37oC and removal of interferon-γ from culture medium. Cells were differentiated for 10-
14 days prior to experiments. The day before imaging, mouse podocytes were infected with adenoviral vectors encoding genes of interest.
Fluorescence microscopy and live cell imaging
For live-cell imaging, Myo1e-null podocytes were plated onto glass-bottom
MatTek dishes that were pre-coated with Collagen IV (MatTek). 24 hours post- infection, cells were observed using Perkin-Elmer UltraView VoX Spinning Disc
Confocal system mounted on a Nikon Eclipse Ti microscope and equipped with an environmental chamber to maintain cells at 37oC. To observe contact reassembly in a calcium switch assay, podocytes were pre-incubated for 5 minutes in Hank’s balanced salt solution (HBSS) containing 2mM EGTA but without calcium and magnesium to remove calcium and magnesium, then
123 Chapter 3 Myo1e motor domain mutants activity in podocytes transferred into warm (pre-warmed to 37oC) Roswell Park Memorial Institute
(RPMI) medium with 10% FBS and normal calcium content and imaged immediately.
Fluorescence Recovery After Photobleaching (FRAP)
FRAP experiments were performed on Perkin-Elmer UltraView VoX
Spinning Disc Confocal system mounted on a Nikon Eclipse Ti-E microscope and equipped with a 561 nm laser and a 60x/1.49 N.A. PlanApo objective as well as an environmental chamber to maintain cells at 37oC. The whole system is controlled by Volocity software. Myo1e-null immortal podocytes were plated on
Collagen IV coated MatTek dishes at an initial density of 1000 cells/dish. Cells were then transferred to 37oC incubator for differentiation for 14 days before imaging. At day 13, podocytes were infected with adenoviral vectors encoding mCherry-tagged ZO-1 alone (KO) or with GFP-tagged Myo1e wild type (KO+WT) or kidney disease associated Myo1e mutants (KO+T119I or KO+A159P). On the day of imaging, fresh complete Roswell Park Memorial Institute (RPMI) medium was added to the MatTek dishes to remove any viruses. For co-expression, only cells that express both markers were chosen to proceed with FRAP experiments.
In addition, only horizontal junctions were chosen in these FRAP experiments in order to maintain the same sized region of interest (ROI).
The FRAP experiments were performed as previously described (Tyska and Mooseker, 2002) with modifications. Briefly, a ROI of 6.8x4.2 μm2 at the
124 Chapter 3 Myo1e motor domain mutants activity in podocytes junction was selected for bleaching (Fig. 4A). For pre-bleaching, initial fluorescence intensity was measured with 5 time points at a rate of 1 sec per time point using 71% laser power and 500 ms exposures. Photobleaching was performed with 100% laser power and scanning the ROI 25 times. Immediately after the bleaching the entire field was scanned at 71% laser power with 500 ms exposures. The first image was taken as t=0. For the first 20 seconds the scan rate was every 4 seconds, and after the initial 20 seconds, the scan rate was reduced to every 10 seconds to prevent further photobleaching. The total imaging time is 300 seconds. The FRAP analysis was performed using ImageJ software
(National Institute of Health, Bethesda, MD) as described in the FRAP manual
(Miura, 2005) and in a previous study (Deakin et al., 2012). The intensity values were normalized such that the scan immediately following the bleach was set to 0 and all subsequent ROI intensity values were expressed relative to this after- bleach value (Deakin et al., 2012). Normalized recovery data was fitted into a double exponential curve using the following equation in KaleidaGraph (Synergy
Software) based on Tyska and Mooseker’s study (Tyska and Mooseker, 2002) with minor modifications:
Y=m2-m3 * exp (-m1 * m0)-m4 * exp (-m5 * m0);
m1=0.5; m2=0.9; m3=0.5; m4=0.1; m5=0.01,
Y is the ROI intensity at time t >=0, m2 is the mobile fraction, m3 is the amplitude of the exponential process with rate k1 (m1) and is equal to the m2 in the
125 Chapter 3 Myo1e motor domain mutants activity in podocytes absence of the second process, and m0 is time. M4 and m5 are the amplitude and rate k2 of the second process, respectively.
All mobile fractions were calculated from the curve, and then averaged and plotted using KaleidaGraph (Synergy Software). Only the total mobile fractions (m2) are used for generating the plot.
Image analysis
All fluorescence intensity measurements were performed with ImageJ software (NIH, Bethesda, MD). Quantitative analysis was performed using
KaleidaGraph (Synergy Software). The statistical significance was calculated using unpaired Student t-test with unequal variance.
126 Chapter 3 Myo1e motor domain mutants activity in podocytes
3.5 Results
3.5.1 The kidney disease associated Myo1e mutants do not co-localize with ZO-
1 at the podocyte junctions.
Myo1e-null podocytes (clone # 1-16) were plated on MatTek dishes and infected with Adenoviral vector encoding GFP-tagged wild type Myo1e or the kidney disease associated Myo1e mutants (Myo1-T119I or A159P). Confocal images show the co-localizations of wild type Myo1e and ZO-1 at cell-cell junctions (Fig. 3.1A-A’’, yellow arrows). The T119I Myo1e mutant does not localize to the junctions (Fig. 3.1B) nor does the Myo1e-A159P as previously noted (Bi et al., 2013). Both of the mutants exhibit diffuse localization in the cytoplasm and lack any specific localization pattern. ZO-1 localization to the cell- cell junctions is not affected by the lack of Myo1e nor by expression of either of the mutants (See Supplemental Fig S2.1 in Chapter 2 and Fig. 3.1B’-B’’ and C’-
C’’, red arrows).
3.5.2 T119I mutant of Myo1e is not recruited to the junctions during calcium switch assay.
In order to determine whether Myo1e T119I mutant affects junction remodeling, we used calcium switch assay that was described previously in chapter 2 (Bi et al., 2013). We infected Myo1e-null podocytes with adenoviral vectors encoding GFP-tagged Myo1e wild type or T119I mutant, and mCherry- tagged ZO-1. Using live-cell imaging, we were able to capture the junction
127 Chapter 3 Myo1e motor domain mutants activity in podocytes reassembly after Ca2+ replenishment. Wild-type Myo1e was recruited to the nascent cell-cell contact at the same time as ZO-1 (Fig. 3.2, yellow arrows).
However, the Myo1e T119I mutant was not recruited to the cell-cell junctions after the Ca2+ replenishment (Fig. 3.3), while the recruitment of ZO-1 to the nascent contact was not altered (Fig. 3.3, red arrows). This is consistent with the other FSGS-associated Myo1e mutant A159P, which is also not recruited to the nascent cell-cell contact (Bi et al., 2013). Interestingly, there was a small amount of mutant Myo1e recruited to the junctions at the beginning of the junction assembly (Fig. 3.3A’, green arrow), which was very similar to the phenotype we saw in our A159P mutant and actin during junction assembly (See Fig. 2.4,
Chapter 2). However, the mutant Myo1e does not co-localize with ZO-1 during junction assembly, revealed by kymograph (Fig. 3.3J).
3.5.3 FRAP analysis reveals faster turnover of mCherry-ZO-1 in cells expressing Myo1e mutants.
To further investigate whether ZO-1 dynamics at the junctions is affected by the different Myo1e motor mutants, we performed FRAP experiments. FRAP analysis is able to measure the lateral mobility of fluorescently labeled molecules.
This includes the half-time of the recovery of the bleached fluorescent molecules and the fraction of these fluorescent molecules that participate in the recovery process (also known as mobile fraction) (Axelrod et al., 1976). Using our Myo1e- null immortal podocytes, we were able to re-introduce GFP-tagged Myo1e wild type or either of the two kidney-disease associated Myo1e motor domain mutants
128 Chapter 3 Myo1e motor domain mutants activity in podocytes
T119I and A159P with mCherry-tagged ZO-1 (Fig. 3.4A). The first 20 sec of the
ZO-1 exchange was very rapid both seen from the live-cell imaging and from the fitted curve. Therefore, we hypothesized that in the first 20 seconds, the ZO-1 exchange was likely due to diffusion. The exchange of the ZO-1 molecule happened later in the process. The ZO-1 fluorescence recovery data was fit best to a double exponential curve, indicating that there are two distinct mechanisms for ZO-1 exchange, one is simple diffusion and the other may result from active transport.
Re-introduction of the kidney disease associated Myo1e motor domain mutant Myo1e-A159P to the Myo1e-null cells increased ZO-1 total mobile fraction
(Fig. 3.4B). However, due to the high variability of the morphology of the podocytes in our cell lines and transient expression level of fluorescently tagged proteins, the ZO-1 mobile fraction was only significantly increased in cells expressing the A159P Myo1e mutant (asterisk in Fig. 3.4B) and not in cells lacking Myo1e or cells expressing T119I mutant. An increase in the ZO-1 mobile fraction in Myo1e-A159P expressing cells suggests that the junctions become more dynamic in these cells and in the case of A159P Myo1e this is likely due to its loss of function as an actin associated protein.
129 Chapter 3 Myo1e motor domain mutants activity in podocytes
3.6 Discussion
The kidney disease-associated Myo1e motor domain mutations, T119I and A159P, disrupt Myo1e localization to the cell-cell junctions, endocytic vesicles, and plasma membranes. Our previous study using fission yeast as a model system showed that these two kidney disease associated Myo1e mutants are non-functional, resulting in growth and endocytosis defects similar to those observed in fission yeast cells lacking Myo1 (Bi et al., 2015). To further explore how these Myo1e motor domain mutants affect podocyte functions, we focused on ZO-1 to study junctional dynamics in kidney podocytes.
Our calcium switch assay suggested that without functional Myo1e, ZO-1 was still recruited to the nascent cell-cell contacts. However, we are unable to conclude whether the efficiency of the ZO-1 recruitment is affected at this time due to lack of statistical analysis. A previous study from another group has used immunostaining and line scan analysis to quantify junctional ZO-1 based on intensity value at the steady state (Maiers et al., 2013). Future investigations to determine whether the junction integrity is affected by expressing of the Myo1e mutants may be performed in the same fashion, using line scan analysis on junctional ZO-1 at different time points during junction assembly. If Myo1e is involved in efficiently recruiting ZO-1 to the junctions, we would expect to see a difference in time or quality of junction assembly in Myo1e mutants expressing cells. In addition, in Chapter 2, we showed that in mutant Myo1e expressing podocytes, the actin assembly at the junctions is defective, which could also be
130 Chapter 3 Myo1e motor domain mutants activity in podocytes linked to ZO-1 recruitment to the junctions, since ZO-1 interacts with actin via its
C-terminus. To determine if Myo1e affects the actin assembly at the junctions, we can perform G-actin corporation assay using fluorescently labeled G-actin
(Kovacs et al., 2011).
To investigate the dynamics of ZO-1 at the steady state, we performed
FRAP experiments to better understand the turnover rate of ZO-1 at the junctions. Our results showed that the fluorescence recovery of ZO-1 in wild type
Myo1e expressing podocytes was about 70%, which was similar to previous studies in other epithelial cell types (Foote et al., 2013; Shen et al., 2008).
According to our recently published paper that both kidney disease associated mutants are non-functional and they have the same loss of function effects on fission yeast cells as Myo1-null strain (Bi et al., 2015), we predict that by expressing the Myo1e-T119I mutant or just using Myo1e-null cells, ZO-1 mobile fractions would also increase in cultured podocytes. However, many more replicates would be required to find the signal in the noise, because our podocyte cell lines are highly variable morphologically. In the future, we can try to use a well-characterized epithelial cell line, such as MDCK cells, to study ZO-1 dynamics at the junctions under the influence of Myo1e mutants.
According to Shen et al., ZO-1 recovery at the junction is mainly due to the exchange of ZO-1 with the intracellular pools (Shen et al., 2008). It would be interesting to further explore which protein, or protein complex, brings the intracellular pool of ZO-1 to the junctions. Shen et al. have also noted that a small
131 Chapter 3 Myo1e motor domain mutants activity in podocytes stable fraction of ZO-1 remains associated with the junctions. We hypothesize that the recovery of this stable fraction of ZO-1 is due to an anchoring molecule or protein complex at the junctions holding ZO-1 in place. If the anchoring molecule is missing or non-functional, the junctions would become less stable and more dynamic. ZO-1 exchange would in turn increase, resulting in a higher mobile fraction.
Increased ZO-1 mobile fraction, especially in A159P expressing podocytes, indicated that the junctions in Myo1e mutants expressing podocytes became more dynamic and less stable. Our early FRAP experiments using podocytes with shRNA knocking down endogenous Myo1e also showed an increased mobile fraction of junctional ZO-1 compared to wild type podocytes
(data not shown). This supports our current model that without functional Myo1e at the junctions, the junctions become less stable, and ZO-1 exchange will increase.
132 Chapter 3 Myo1e motor domain mutants activity in podocytes
3.7 Reference:
Al-Hamed, M.H., Al-Sabban, E., Al-Mojalli, H., Al-Harbi, N., Faqeih, E., Al Shaya, H., Alhasan, K., Al-Hissi, S., Rajab, M., Edwards, N., Al-Abbad, A., Al-Hassoun, I., Sayer, J. a, Meyer, B.F., 2013. A molecular genetic analysis of childhood nephrotic syndrome in a cohort of Saudi Arabian families. J. Hum. Genet. 1–10. doi:10.1038/jhg.2013.27 Axelrod, D., Koppel, D.E., Schlessinger, J., Elson, E., Webb, W.W., 1976. Mobility measurement by analysis of fluorescence photobleaching recovery kinetics. Biophys. J. 16, 1055–1069. doi:10.1016/0002-9610(76)90409-8 Bi, J., Carroll, R.T., James, M.L., Ouderkirk, J.L., Krendel, M., Sirotkin, V., 2015. Effects of FSGS-associated mutations on the stability and function of myosin-1 in fission yeast. Dis. Model. Mech. 8(8):891-902. doi:10.1242/dmm.020214 Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M., 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am. J. Physiol. Renal Physiol. 305, 532–544. doi:10.1152/ajprenal.00223.2013 D’Agati, V.D., Kaskel, F.J., Falk, R.J., 2011. Focal segmental glomerulosclerosis. N. Engl. J. Med. 365, 2398–411. doi:10.1056/NEJMra1106556 Deakin, N.O., Ballestrem, C., Turner, C.E., 2012. Paxillin and Hic-5 interaction with vinculin is differentially regulated by Rac1 and RhoA. PLoS One 7. doi:10.1371/journal.pone.0037990 Faul, C., Asanuma, K., Yanagida-Asanuma, E., Kim, K., Mundel, P., 2007. Actin up: regulation of podocyte structure and function by components of the actin cytoskeleton. Trends Cell Biol. 17, 428–37. doi:10.1016/j.tcb.2007.06.006 Foote, H.P., Sumigray, K.D., Lechler, T., 2013. FRAP analysis reveals stabilization of adhesion structures in the epidermis compared to cultured keratinocytes. PLoS One 8, e71491. doi:10.1371/journal.pone.0071491 Fukasawa, H., Bornheimer, S., Kudlicka, K., Farquhar, M.G., 2009. Slit diaphragms contain tight junction proteins. J. Am. Soc. Nephrol. 20, 1491–503. doi:10.1681/ASN.2008101117 Jat, P.S., Noble, M.D., Ataliotis, P., Tanaka, Y., Yannoutsos, N., Larsen, L., Kioussis, D., 1991. Direct derivation of conditionally immortal cell lines from an H-2Kb-tsA58 transgenic mouse. Proc. Natl. Acad. Sci. U. S. A. 88, 5096–5100. doi:10.1073/pnas.88.12.5096 Kim, S. V, Mehal, W.Z., Dong, X., Heinrich, V., Pypaert, M., Mellman, I., Dembo, M., Mooseker, M.S., Wu, D., Flavell, R. a, 2006. Modulation of cell adhesion and motility in the immune system by Myo1f. Science 314, 136–9. doi:10.1126/science.1131920 Kovacs, E.M., Verma, S., Ali, R.G., Ratheesh, A., Hamilton, N. a., Akhmanova, A., Yap, A.S., 2011. N-WASP regulates the epithelial junctional actin cytoskeleton through a non- canonical post-nucleation pathway. Nat. Cell Biol. 13, 934–943. doi:10.1038/ncb2290 Krendel, M., Kim, S. V, Willinger, T., Wang, T., Kashgarian, M., Flavell, R. a, Mooseker, M.S., 2009. Disruption of Myosin 1e promotes podocyte injury. J. Am. Soc. Nephrol. 20, 86–94. doi:10.1681/ASN.2007111172 Maiers, J.L., Peng, X., Fanning, A.S., DeMali, K. a, 2013. ZO-1 recruitment to α- catenin--a novel mechanism for coupling the assembly of tight junctions to adherens junctions. J. Cell Sci. 126, 3904–15. doi:10.1242/jcs.126565 Mathieson, P.W., 2011. The podocyte as a target for therapies-new and old. Nat. Rev. Nephrol. 1–5. doi:10.1038/nrneph.2011.171 Mele, C., D, B.S., Iatropoulos, P., Donadelli, R., Calabria, A., Eng, D., Maranta, R., Cassis, P., Ph, D., Buelli, S., Tomasoni, S., Piras, R., D, C.P., Krendel, M., Bettoni, S., Biotech, D., Morigi, M., Delledonne, M., Pecoraro, C., Abbate, I., Capobianchi, M.R., Hildebrandt, F., Otto, E., Schaefer, F., Macciardi, F., Ozaltin, F., Emre, S., Ibsirlioglu, T., Benigni, A., Remuzzi, G., 2011. MYO1E mutations and childhood familial focal segmental glomerulosclerosis. N. Engl. J. Med. 365, 295–306. Miura, K., 2005. Analysis of FRAP Curves. EMBL 1–35.
133 Chapter 3 Myo1e motor domain mutants activity in podocytes
Mundel, P., Reiser, J., Zúñiga Mejía Borja, a, Pavenstädt, H., Davidson, G.R., Kriz, W., Zeller, R., 1997. Rearrangements of the cytoskeleton and cell contacts induce process formation during differentiation of conditionally immortalized mouse podocyte cell lines. Exp. Cell Res. 236, 248–58. Reiser, J., Kriz, W., Kretzler, M., Mundel, P., 2000. The glomerular slit diaphragm is a modified adherens junction. J. Am. Soc. Nephrol. 11, 1–8. Rood, I.M., Deegens, J.K.J., Wetzels, J.F.M., 2012. Genetic causes of focal segmental glomerulosclerosis: implications for clinical practice. Nephrol. Dial. Transplant 27, 882–90. doi:10.1093/ndt/gfr771 Sanna-Cherchi, S., Burgess, K.E., Nees, S.N., Caridi, G., Weng, P.L., Dagnino, M., Bodria, M., Carrea, A., Allegretta, M. a, Kim, H.R., Perry, B.J., Gigante, M., Clark, L.N., Kisselev, S., Cusi, D., Gesualdo, L., Allegri, L., Scolari, F., D’Agati, V., Shapiro, L.S., Pecoraro, C., Palomero, T., Ghiggeri, G.M., Gharavi, A.G., 2011. Exome sequencing identified MYO1E and NEIL1 as candidate genes for human autosomal recessive steroid-resistant nephrotic syndrome. Kidney Int. 80, 389–96. doi:10.1038/ki.2011.148 Shen, L., Weber, C.R., Turner, J.R., 2008. The tight junction protein complex undergoes rapid and continuous molecular remodeling at steady state. J. Cell Biol. 181, 683–695. doi:10.1083/jcb.200711165 Tyska, M.J., Mooseker, M.S., 2002. MYO1A (brush border myosin I) dynamics in the brush border of LLC-PK1-CL4 cells. Biophys. J. 82, 1869–1883. doi:10.1016/S0006- 3495(02)75537-9 Welsh, G.I., Saleem, M. a, 2011. The podocyte cytoskeleton-key to a functioning glomerulus in health and disease. Nat. Rev. Nephrol. 1–8. doi:10.1038/nrneph.2011.151
134 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.1. Kidney disease associated Myo1e mutants do no co-localize with ZO-1 at the cell-cell junctions.
135 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.1. Kidney disease associated Myo1e mutants do no co-localize with ZO-1 at the cell-cell junctions. Adenoviral vectors encoding GFP-tagged
Myo1e wild type (A) or mutants (B-C), and mCherry-tagged ZO-1 (A’-C’) were infected to Myo1e-null mouse podocytes. The merged images are shown in A’’-
C’’. Yellow arrows in A indicate the co-localization of wild type Myo1e and ZO-1 at the junctions, while red arrows in B and C point to ZO-1 localization at the junctions without colocalization with the Myo1e mutants T119I and A159P. Scale bar: 10 μm.
136 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.2. Myo1e is recruited to the nascent cell-cell contact at the same time as ZO-1 during the junction assembly.
137 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.2. Myo1e is recruited to the nascent cell-cell contact at the same time as ZO-1 during the junction assembly. Myo1e-null mouse podocytes were infected with GFP-tagged wild type Myo1e (A, D, and G) and mCherry- tagged ZO-1 (B, E, F). Individual frames from a time-lapse video of the calcium switch experiment at the time points of 1 min (A-C), 4 min (D-F) and 8 min (G-I) are shown. The zoomed-in images from the boxed areas in images A, D, and G are displayed in the right-hand panels (A’-F’). Yellow arrows indicate the recruitment and co-localization of wild type Myo1e and ZO-1. Scale bar: 10 μm.
138 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.3. Kidney disease-associated Myo1e mutant T119I is not recruited to the nascent cell-cell junction with ZO-1 at the same time.
139 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.3. Kidney disease-associated Myo1e mutant T119I is not recruited to the nascent cell-cell junction with ZO-1 at the same time. Myo1e-null mouse podocytes were infected with GFP-tagged mutant Myo1e-T119I (A, D, and G) and mCherry-tagged ZO-1 (B, E, F). Individual frames from a time-lapse video of the calcium switch experience at the time point of 1 min (A-C), 5 min (D-
F) and 10 min (G-I) are shown. The zoomed-in images from the boxed areas in images A, D, and G are displayed in the right-hand panels (A’-F’). Red arrows indicate the recruitment of ZO-1 only at the nascent cell-cell contact during calcium switch experiment at a certain time point. The green arrow in A’ refers to the recruitment of Myo1e-T119I at the beginning of junction assembly, which does not co-localize with ZO-1, indicated by Kymograph (J). Scale bar: 10 μm.
140 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.4. ZO-1 mobile fractions are increased in cells lacking Myo1e or expressing kidney disease-associated Myo1e mutants.
141 Chapter 3 Myo1e motor domain mutants activity in podocytes
Figure 3.4. ZO-1 mobile fractions are increased in cells lacking Myo1e or expressing kidney disease-associated Myo1e mutants. (A) Fluorescence
Recovery After Photobleaching (FRAP) experiment is demonstrated. The yellow- boxed region at the junction between cell 1 and cell 2 indicates the bleached region. Before photobleaching (pre-bleach), the fluorescence intensity inside the boxed region is recorded as baseline and normalized to 1. After the photobleaching (post-bleach), the fluorescence recovery inside the boxed region was recorded at a certain frequency (T1, T2, …). The data was analyzed and normalized, and plotted as double exponential curve (detailed information see
Material and Methods). (B) Bar graphs show that the mobile fraction of ZO-1 is higher in the Myo1e-null podocytes or cells expressing Myo1e mutants T119I or
A159P. Asterisk (*) indicates the data is statistically significant. P=0.062 (KO),
0.0024 (KO+Myo1eA159P), 0.075 (KO+Myo1eT119I).
142 Chapter 4 Myo1e mutants in yeast
Chapter 4
Effects of FSGS-associated mutations on the
stability and function
of myosin-1 in fission yeast
Chapter 4 is adapted from the following article published in 2015: Bi, J., Carroll, R.T., James, M.L., Ouderkirk, J.L., Krendel, M., Sirotkin, V., 2015. Effects of FSGS- associated mutations on the stability and function of myosin-1 in fission yeast. Dis. Model. Mech., 8(8):891-902.
143 Chapter 4 Myo1e mutants in yeast
4.1 Attributions and Acknowledgments
J.B., R.T.C., M.K., and V.S. designed the experiments. J.B., R.T.C., M.J., and V.S. performed the experiments. J.L.O. prepared the myo1-A181P construct.
J.B. analyzed the data and prepared Figures. J.B., M.K., and V.S. wrote the manuscript with input from R.T.C and M.J.
We thank Dr. Matthew Lord (University of Vermont) for yeast strains. We are grateful to the members of Krendel, Sirotkin, Pruyne and Amack labs (SUNY
Upstate Medical University) for their input and critical discussions of this project.
We thank out funding source American Heart association (AHA Predoctoral
Fellowship grant 14PRE20380534 to J. Bi, and SDG 11SDG5470024 to V.
Sirotkin), National Institute of Health and NIDDK (R01DK083345 to M. Krendel)
144 Chapter 4 Myo1e mutants in yeast
4.2 Abstract
Point mutations in the human MYO1E gene, encoding class I myosin
Myo1e, are associated with focal segmental glomerulosclerosis (FSGS), a primary kidney disorder that leads to end stage kidney disease. In this study, we used a simple model organism, fission yeast Schizosaccharomyces pombe, to test the effects of FSGS-associated mutations on myosin activity. Fission yeast has only one class I myosin, Myo1, which is involved in actin patch assembly at the sites of endocytosis. The amino acid residues mutated in the FSGS patients are conserved between human Myo1e and yeast Myo1, which allowed us to introduce equivalent mutations into yeast myosin and use the resulting mutant strains for functional analysis. Yeast strains expressing mutant Myo1 exhibited defects in growth and endocytosis similar to those observed in the myo1 deletion strain. These mutations also disrupted Myo1 localization to endocytic actin patches and resulted in mis-localization of Myo1 to eisosomes, linear membrane microdomains found in yeast cells. While both mutants examined in this study exhibited loss of function, one of these mutants was also characterized by the decreased protein stability. Thus, using the yeast model system we were able to determine that the kidney disease-associated mutations impair myosin functional activity and have differential effects on protein stability.
145 Chapter 4 Myo1e mutants in yeast
4.3 Introduction
Missense mutations in the human MYO1E gene encoding molecular motor protein Myosin 1e (Myo1e) are associated with kidney disease (Al-Hamed et al.,
2013; Mele et al., 2011; Sanna-Cherchi et al., 2011). While these mutations have been predicted to disrupt Myosin 1e activity, testing this prediction directly in a vertebrate model organism is challenging, not only due to the need for complex genetic modifications but also because of the limitations of microscopic and biochemical characterization in such a model. Therefore, we took advantage of a favorable model system, fission yeast S. pombe, to examine the effects of disease-associated mutations on myosin-1 localization, activity, and stability.
From more than twenty known classes of myosins, approximately forty myosins distributed among 12 classes are present in humans (Berg et al., 2001), including 8 members of class I myosins. Several myosins have been implicated in human health and disease (Moore et al., 2012; Ouderkirk and Krendel, 2014;
Redowicz, 2002; Tajsharghi and Oldfors, 2013). Mutations in the MYO1E gene in humans are associated with focal segmental glomerulosclerosis (FSGS) (Al-
Hamed et al., 2013; Mele et al., 2011; Sanna-Cherchi et al., 2011), a kidney disease arising from the disruption of the protein filtration barrier in the kidney, which leads to proteinuria (protein excretion in the urine). FSGS is characterized by the structural changes in the renal glomeruli, the portions of the nephrons that are responsible for selective protein filtration. In many cases of familial FSGS, the disruption of renal filtration arises from the defects in specialized epithelial cells of
146 Chapter 4 Myo1e mutants in yeast the glomerulus, called podocytes (Greka and Mundel, 2012). Podocytes have long, actin-rich, interdigitating cell processes, called foot processes, which cover the entire surface of the glomerular capillaries. The contacts between podocyte foot processes, also known as the slit diaphragms, represent one of the key components of the glomerular filtration barrier (George and Holzman, 2012;
Greka and Mundel, 2012).
Myosin 1e (Myo1e) is expressed in podocytes and plays a key role in regulating the integrity of their cell-cell junctions (Bi et al., 2013). Myo1e-null mice develop proteinuria at 2 weeks after birth, along with the structural changes in the glomeruli that are reminiscent of those observed in patients with FSGS (Krendel et al., 2009; Mele et al., 2011). While it is known that the complete knockout of
Myo1e or its selective knockout in podocytes are sufficient to induce FSGS-like disease in mice (Chase et al., 2012; Krendel et al., 2009), the effects of substituting wild-type Myo1e with the mutant versions associated with human
FSGS have not been tested in a model organism.
Like other members of the myosin superfamily, human Myo1e and S. pombe class I myosin Myo1 have a highly conserved N-terminal motor domain
(Fig. 4.1A) that uses ATP hydrolysis to move along an actin filament and a C- terminal tail domain that binds membrane and protein cargo (Bement et al., 1994;
El Mezgueldi et al., 2002; Feeser et al., 2010; Krendel et al., 2007; Lee et al.,
2000; Sirotkin et al., 2005). Both missense mutations in the MYO1E gene identified in the FSGS patients result in changes in conserved amino acid
147 Chapter 4 Myo1e mutants in yeast residues in the myosin motor domain. The T119I mutation (Al-Hamed et al.,
2013) is located in the P-loop region, while the A159P mutation (Mele et al.,
2011; Sanna-Cherchi et al., 2011) is located adjacent to the Switch-I region (Fig.
4.1B). These regions are involved in ATP binding and hydrolysis. Therefore, mutations in these highly conserved residues are likely to disrupt the functions of the myosin motor domain.
In order to determine whether a particular disease-associated mutation is pathogenic or represents a non-pathogenic polymorphism, it is important to test the effects of each mutation on protein functions. While the effects of T119I on myosin-1 function are yet to be tested, we have previously characterized the effects of the A159P mutation on Myo1e activity by expressing the mutant Myo1e in the Myo1e-null podocytes (Bi et al., 2013). The mutant was mis-localized, and cells expressing this Myo1e variant exhibited defects in cell-cell contact formation. However, this analysis was complicated by the inherent variability of contact assembly between individual pairs of cells detected by live cell imaging and by the lack of clear understanding of Myo1e function in contact assembly.
In the present study, we tested the effects of mutations equivalent to a known Myo1e pathogenic mutation (A159P) and a novel mutation recently identified by candidate gene sequencing (T119I) (Al-Hamed et al., 2013) in a favorable model system S. pombe. Unlike vertebrates that have multiple class I myosins, fission yeast have a single class 1 myosin, Myo1, and its localization and function are well-defined. Myo1 assembles into actin patches at the sites of
148 Chapter 4 Myo1e mutants in yeast endocytosis in an ordered, highly reproducible manner and plays a role in promoting endocytic internalization (Sirotkin et al., 2010). Yeast lacking Myo1 exhibit growth defects, abnormal morphology, and changes in actin patch localization and dynamics (Lee et al., 2000; Sirotkin et al., 2005). We took advantage of the conservation of motor domains between Myo1e and Myo1 to examine the effects of the FSGS-equivalent mutations on fission yeast Myo1.
Intriguingly, we found that while both mutations disrupted Myo1 localization and function, they had differential effects on Myo1 stability and folding as gauged by immunoblotting and by co-localization with a myosin chaperone Rng3 (Lord and
Pollard, 2004; Stark et al., 2013).
149 Chapter 4 Myo1e mutants in yeast
4.4 Materials and Methods
Yeast strains
Yeast strains used in this study are listed in Supplementary Table S1. All strains were constructed by genomic integrations using homologous recombination (Bahler et al., 1998; Sammons et al., 2011) and genetic crosses.
The myo1-E1 (G308R) and myo1-S1 (G483D) mutants were generated previously (Stark et al., 2013) based on the sequence homology with myo2-E1 and myo2-S1 mutants (Balasubramanian et al., 1998; Wong et al., 2000).
Nucleotide changes corresponding to the T140I and A181P mutations were introduced into S. pombe myo1 gene sequence using QuickChange kit
(Stratagene; La Jolla, California) according to the manufacturer’s instructions.
The mutations were generated in the pBS-myo1 construct that contains 5 kb
EcoR1 fragment of S. pombe genomic DNA encompassing myo1 gene (Lee et al., 2000). To integrate these mutations into the endogenous myo1 locus, mutated constructs were digested with EcoR1, and 5 kb myo1 gene fragments were introduced via lithium acetate transformation into myo1Δ::ura4+ strain
(TP192), in which myo1 open reading frame is replaced with the ura4+ nutritional marker (Sirotkin et al., 2005), followed by counter-selection against ura4+ on
EMM plates containing 2 mg/ml 5-FOA and 0.05 mg/ml uracil. Successful integrations of mutated myo1 sequences were confirmed by sequencing of PCR- amplified fragments of myo1 locus.
150 Chapter 4 Myo1e mutants in yeast
To tag wild type and mutant Myo1 with mGFP or mCherry at the C- terminus, the stop codon in the endogenous myo1 gene in normal chromosomal location was replaced with gene tagging cassettes PCR-amplified from pFA6a- mGFP-kanMX6 (pJQW 85-4) for mGFP (Bahler et al., 1998; Wu et al., 2003), or pFA6a-mCherry-natMX6 (pKS391) for mCherry (Snaith et al., 2005).
Haploid strains combining mutants, mGFP- or mCherry-tagged markers were generated by genetic crosses. Parental strains were crossed on ME plates followed by tetrad dissection directly from the cross or from the sporulating diploids isolated from the crosses. For some crosses, G308R mutant was transformed with pUR-myo1+ plasmid, which facilitates crosses and is lost upon sporulation. During constructing the strains, no effects of tagging Myo1 mutants with fluorescent protein tags or combining these mutants with fluorescent protein- tagged Fim1, Cam2, Lifeact, or Rng3 on cell viability were observed.
Fluorescence microscopy
Fission yeast cells were grown at 25oC in YE5S medium to exponential phase. Cells were then washed in EMM medium and mounted on pads of 25% gelatin in EMM under coverslips. The images were taken using PerkinElmer
UltraView VoX Spinning Disc Confocal system mounted on a Nikon Eclipse Ti-E microscope equipped with Hamamatsu C9100-50 EMCCD camera and a
100x/1.4 N.A. PlanApo objective and controlled by Volocity software. The Z- series through the entire cells were captured at 0.4-µm intervals, and the time- lapse images in a single confocal section in the middle plane of the cell were
151 Chapter 4 Myo1e mutants in yeast collected at a 2-second intervals for 1 min. Maximum intensity projections were generated from three Z-sections through the top surface of the cells.
Image analysis
All image analysis was performed in ImageJ (National Institutes of Health,
Bethesda, MD). Brightness and contrast were adjusted identically within each figure. For actin patch tracking, at least 5 patches were selected from a time- lapse movie for each strain. The patch fluorescence intensity, position, and movement were measured in ImageJ. Time courses of intensity and distance from origin for individual patches were aligned in time to peak intensity of Fim1- mCherry (time zero) and averaged at each time point. The whole cell intensities in GFP (Fig. 1G) channel were measured for 5 cells in frame 4 of the time series in a single confocal section through the middle of the cells and subtracted for extracellular background. Co-localization between mGFP-tagged Myo1 and mCherry-tagged Fimbrin (Fig. S2A) was measured in ImageJ by selecting all
Fim1-mCherry patches at frame 10 and counting the fraction of Fim1-mCherry patches that contained mGFP-labeled Myo1 within the 10 pixel selection area centered on Fim1-mCherry patch at any time point during Fim1-mCherry patch lifetime. The average numbers of eisosome per surface area (Fig. S3) were measured by counting the numbers of eisosomes in the maximum intensity projections of three Z-sections through the top surface of three cells. Co- localization between Myo1 and Cam2 in patches, puncta, and eisosomes (Fig.
S5) were measured from three Z-sections through the top surface of three cells.
152 Chapter 4 Myo1e mutants in yeast
For patches, the presence of both mGFP and mCherry signals within 10-pixel wide selection area centered on a patch was counted as co-localization.
Western blots
Fission yeast cells were grown under the same conditions that were used for live cell imaging. Cells were harvested by centrifugation and processed for immunoblotting (Wu and Pollard, 2005). Pelleted cells were re-suspended in ice- cold lysis buffer U (50mM HEPES pH7.5, 100mM KCl, 3mM MgCl2, 1mM EGTA,
1mM EDTA, 0.1% TritonX-100, 1mM DTT, 1mM PMSF, and Complete (Roche,
NJ) protease inhibitor cocktail). Total protein samples were prepared by mechanical lysis with glass beads in FastPrep-24 (MP-Bio, Santa Ana,
California), followed by addition of SDS-PAGE sample buffer, 2 min incubation at
100oC and 2 min centrifugation at 14,000 rpm in a microcentrifuge. Samples containing equal amounts of total protein, adjusted based on quantitation of
Coomassie-stained gels, were separated on 10-20% gradient SDS-PAGE gels and transferred to Immobilon-P (EMD Millipore, Billerica, MA) membranes followed by incubation with 1:3000 dilution of primary anti-GFP antibody (Pierce,
Rockford, IL) overnight at 4oC and secondary horseradish peroxidase-conjugated antibody for 1 hr at room temperature. Blots were developed using Clarity™
Western ECL substrate (BioRad, Hercules, CA) and imaged using Bio-Rad
(BioRad, Hercules, CA) ChemiDoc MP imager.
The total and the full length band intensities were measured in Image Lab
(BioRad, Hercules, CA) and normalized to the wild type (Fig. 1D-F). Fraction of
153 Chapter 4 Myo1e mutants in yeast the total band intensity detected in the full-length band was also calculated (Fig.
S1A).
Cycloheximide treatment
Cycloheximide (CHX) treatment was performed as previously described
(Roseaulin et al., 2013). Briefly, fission yeast cells were grown at 25oC in YE5S medium to exponential phase. 0.1 mg/ml CHX (Amresco, Solon, Ohio) were added to cells for 24hrs at 25oC, and cells were lysed and processed for western blot analysis.
154 Chapter 4 Myo1e mutants in yeast
4.5 Results
4.5.1 Mutations in the motor domain of Myo1, equivalent to the FSGS- associated Myo1e mutations, result in yeast growth defects
Based on sequence alignment (Fig. 4.1B), we identified conserved residues T140 and A181 in the S. pombe Myo1 motor domain as homologous to the residues T119 and A159 in human Myo1e, which are disrupted by the T119I and A159P mutations in human FSGS patients. To examine the effects of these mutations in fission yeast, we replaced the wild type myo1+ coding sequence with the mutant myo1 variants directly in the yeast genome so that the mutant myosins were expressed from the myo1 genomic locus under the control of the endogenous promoter. Like Δmyo1 (Lee et al., 2000), myo1(T140I) and myo1(A181P) cells were viable and were able to grow on rich YES medium at
25oC (Fig. 4.1C).
Many endocytosis mutants in budding and fission yeast, including Δmyo1
(Lee et al., 2000; Munn et al., 1995; Whitacre et al., 2001), exhibit sensitivity to high temperature (36oC) and high salt (1M KCl or NaCl). To examine the effects of Myo1 motor domain mutations on cell growth, we directly compared the ability of myo1(T140I), myo1(A181P), Δmyo1 and wild type cells to grow on rich YES medium containing 1M KCl at 36oC and on minimal EMM medium containing
Phloxine B (PB), an indicator of cell viability, at 25oC and 36oC (Fig. 4.1C).
Increased PB staining indicates decreased viability. Wild type cells grew on
YE5S+1M KCl plates and stained lightly pink on EMM+PB plates at 25oC and
155 Chapter 4 Myo1e mutants in yeast
36oC. In contrast, myo1(T140I) and myo1(A181P) cells completely failed to grow on YES+1M KCl plates and grew but stained dark red on EMM+PB plates at both
25oC and 36oC, indicating decreased viability. Salt sensitivity and decreased viability of myo1(T140I) and myo1(A181P) cells were identical to those observed in Δmyo1 cells. Thus, each of the two motor domain mutations disrupted Myo1 function in yeast cells to the same degree as complete myo1 deletion.
4.5.2 A181P but not T140I mutation results in the decreased stability of Myo1 in yeast
To examine the effects of motor domain mutations on Myo1 stability and localization in cells, we tagged both mutants with monomeric GFP (mGFP) or mCherry at the C-terminus directly in the native myo1 chromosomal location in haploid cells. The tagged proteins were expressed under the control of endogenous myo1 promoter and were the sole source of Myo1 in cells. We compared the stability and localization of mGFP-tagged proteins to similarly tagged wild type Myo1 and two previously constructed Myo1 motor domain mutants, Myo1-E1 (G308R) and Myo1-S1 (G483D) (Stark et al., 2013). The
Myo1-E1 and Myo1-S1 mutants were modeled after the severe Myo2-E1 and mild Myo2-S1 mutants of S. pombe myosin-II Myo2 (Balasubramanian et al.,
1998; Wong et al., 2000). The G345R mutation in the myo2-E1 allele destabilizes
Myo2, resulting in cytokinesis defects and enhanced recruitment of myosin chaperone Rng3 to the contractile ring, while the G515D mutation in the myo2-S1 allele does not (Stark et al., 2013; Wong et al., 2000).
156 Chapter 4 Myo1e mutants in yeast
Western blot analysis of yeast extracts using anti-GFP antibodies revealed increased degradation of the mGFP-tagged A181P and G308R Myo1 mutants compared to the mGFP-tagged wild type Myo1, T140I and G483D Myo1 mutants
(Fig. 4.1D-F and Fig. S4.1A). Under our lysis conditions, even wild-type Myo1- mGFP was susceptible to degradation such that 30% was detected as the ~150 kDa full-length Myo1-mGFP (arrow in Fig. 4.1D and Fig. S4.1A), and the rest as lower molecular weight degradation products with a prominent ~40 kDa band
(arrowhead in Fig. 4.1D). A similar degradation pattern and similar amounts of the total mGFP-tagged protein were observed for Myo1(T140I)-mGFP and
Myo1(G483D)-mGFP, although compared to the wild type Myo1-mGFP, more
Myo1(T140I)-mGFP and less Myo1(G483D)-mGFP were detected as the full- length protein (Fig. 1D and Fig. S1A). In contrast, the total amounts of
Myo1(A181P)-mGFP and Myo1(G308R)-mGFP were substantially reduced to 0.6 and 0.4 of the wild type level (Fig. 4.1E), respectively, and only 10% of the total was detected as the full length protein (Fig. 4.1F and Fig. S4.1A). Myo1(A181P)- mGFP and Myo1(G308R)-mGFP also showed a different degradation pattern: the amounts of the 40 kDa degradation product were greatly reduced and replaced with a new, lower molecular weight ~30 kDa product (asterisk in Fig. 4.1D), representing almost complete degradation of mutant Myo1. Unlike the level of protein detected by immunoblotting, the total fluorescence intensities of cells before lysis (Fig. 4.1G), representing total intracellular concentrations (Wu and
Pollard, 2005), were the same for all strains, indicating that all mutants were
157 Chapter 4 Myo1e mutants in yeast expressed at the same level, and that the degradation likely occurred in part after cell lysis. Treatment with 0.1 mg/ml cycloheximide for 24 hours to block new protein synthesis had no effect on the degradation pattern and amounts of the detected protein (Fig. S4.1B), suggesting that in cells wild type and mutant Myo1 turn over very slowly and their degradation products, once generated, remain stable in the cytoplasm. Increased degradation detected by immunoblotting suggested decreased stability of A181P and G308R Myo1 mutants. Surprisingly, from the two FSGS-associated mutations, only the A181P mutation decreased
Myo1 stability, whereas the T140I mutation did not.
4.5.3 Myo1 motor domain mutations disrupt localization and function of Myo1 in endocytic actin patches
To characterize the effects of motor domain mutations on Myo1 localization in live fission yeast cells, we examined localization of mGFP-tagged wild type and mutant Myo1 variants relative to the endocytic actin patches labeled with mCherry-tagged actin-binding protein fimbrin Fim1 (Fig. 4.2). While the wild-type Myo1-mGFP and the mild mutant Myo1(G483D)-mGFP co-localized with Fim1-mCherry in more than 95% endocytic actin patches on the cell surface
(Fig. S4.2A and yellow arrowheads in Fig. 4.2A-B’’), the three other motor domain mutants, Myo1(T140I)-mGFP, Myo1(A181P)-mGFP, and Myo1(G308R)- mGFP did not (Fig. 4.2D-F’’). Instead, these three Myo1 motor domain mutants localized to filamentous structures at the cell cortex (green arrows in Fig. 4.2D–
F’’), which had only nominal, less than 10% overlap with actin patches (Fig.
158 Chapter 4 Myo1e mutants in yeast
S4.2A). Thus, functional motor domain is required for Myo1 localization to actin patches. Interestingly, approximately 3-fold more Myo1(T140I)-mGFP localized to these filamentous cortical structures, subsequently identified as eisosomes, than
Myo1(A181P)-mGFP or Myo1(G308R)-mGFP (Fig. S4.2B-E), which correlated with the amount of full-length protein detected by immunoblotting (Fig. 4.1 and
Fig. S4.1). This suggests that the increased degradation of A181P and G308R
Myo1 mutants decreases their localization at the cell cortex.
Since Myo1(T140I)-mGFP, Myo1(A181P)-mGFP, and Myo1(G308R)- mGFP failed to localize to actin patches, we compared actin patch dynamics measured with Fim1-mCherry in Myo1 motor domain mutants, Δmyo1 or wild type cells (Fig. 4.3A-F). In the wild type cells, patches assembled in ~7 seconds to peak intensity corresponding to ~900 fimbrin molecules (Sirotkin et al., 2010) and disassembled in 9 seconds, and nearly all patches moved away from the cortex, representing successful endocytic internalization. The wild type Myo1 appeared 1-2 seconds before Fim1, peaked 1-2 seconds before Fim1, and stayed on the membrane as actin patch associated with the endocytic vesicle internalized (Fig. 4.3A), which is why co-localization between Myo1 and Fim1 is not perfect (Sirotkin et al., 2005). The same actin patch dynamics was observed in cells expressing the mild Myo1(G483D)-mGFP mutant (Fig. 4.3B). In contrast, mGFP-tagged T140I, A181P, and G308R Myo1 mutants completely failed to localize to actin patches at any point during the patch lifetime (Fig. 4.3D-F), and cells expressing these mutants exhibited the same defects in patch assembly
159 Chapter 4 Myo1e mutants in yeast and constitutive endocytic internalization (Fig. 4.3G-I) as in the Δmyo1 cells (Fig.
4.3C). In cells expressing Myo1(T140I)-mGFP, Myo1(A181P)-mGFP, and
Myo1(G308R)-mGFP, patches exhibited a 12-20 seconds longer lifetime (T140I:
32.8+/-5.8 seconds; A181P: 28.0+/-8.4 seconds; G308R: 36.0+/-6.3 seconds) due to 6-8 seconds longer assembly time (T140I: 13.8+/-1.8 seconds; A181P:
12.6+/-4.3 seconds; G308R: 15.4+/-4.3 seconds) and 6-12 seconds longer disassembly time compared to the wild type cells (Fig. 4.3G). While patches in these three motor domain mutants assembled to the similar peak intensity as in the wild type cells (Fig. 4.3I), only half of the patches successfully internalized
(Fig. 4.3H), indicating a significant defect in constitutive endocytosis, which correlated with the reduced viability of these cells (Fig. 4.1C). Thus, motor domain mutations prevent Myo1 localization to patches and result in the same defects of actin patch dynamics as the complete myo1 deletion.
4.5.4 Myo1 motor domain mutants co-localize with the eisosome marker Fhn1
Elongated cortical structures decorated by mGFP-tagged T140I, A181P, and G308R Myo1 motor domain mutants were reminiscent of eisosomes.
Eisosomes are membrane invaginations/microdomains characterized by the presence of specific membrane-binding and transmembrane proteins, including
Fhn1 (Kabeche et al., 2011; Moreira et al., 2009; Olivera-Couto et al., 2011;
Walther et al., 2007). To determine if Myo1 motor domain mutants indeed localized to eisosomes, we combined by genetic crosses mCherry-tagged eisosome marker Fhn1 with mGFP-tagged wild type or mutant Myo1 (Fig. 4.4).
160 Chapter 4 Myo1e mutants in yeast
Fluorescence imaging revealed co-localization of Myo1 mutants T140I, A181P, and G308R with Fhn1 in eisosomes (Fig. 4.4C-E’’, yellow arrows), whereas wild type Myo1 or G483D mutant localized to actin patches (Fig. 4.4A-B’’, green arrowheads) that showed little overlap with Fhn1-mCherry labeled eisosomes
(Fig. 4.4A-B’’, red arrows). Thus, disruption of Myo1 motor domain by kidney disease-associated mutations resulted in targeting of Myo1 to eisosomes. The association of Myo1 mutants with eisosomes appeared to have no effect on eisosome formation since the number of eisosomes in wild type and myo1 mutant cells was approximately the same (Fig. S4.3), despite differences in the amount of Myo1 associated with eisosomes among Myo1 mutants (Fig. S4.2).
To test whether mutant Myo1 in eisosomes can recruit any filamentous actin (F-actin) and, conversely, whether any eisosome-localizing Myo1 mutants can associate with actin patches, cables, and rings, which are the three actin structures in fission yeast cells, we examined the localizations of mGFP-tagged
Myo1 wild type and mutant Myo1 variants relative to the actin structures labeled with a general F-actin marker Lifeact-mCherry (Fig. 4.5). In the wild type cells,
Myo1 co-localized with Lifeact in patches (Fig. 4.5A-A’’’, yellow arrowheads) but not in cables (Fig. 4.5A-A’’’, red arrows) or contractile actin rings. In contrast, two kidney disease associated mutants, Myo1(T140I)-mGFP and Myo1(A181P)- mGFP, localized to eisosomes (Fig. 4.5B-C’’’, green arrows) that were clearly distinct from any Lifeact-labeled F-actin structures in the cells (Fig. 4.5B-C’’’, red arrows, filled and open arrowheads). In the two mutants, as in Dmyo1 cells, all F-
161 Chapter 4 Myo1e mutants in yeast actin structures, including patches, lacked any Myo1 and, conversely, no Lifeact- labeled F-actin was detected associated with Myo1 mutants in eisosomes.
4.5.5 Myo1 mutants co-localize with the Cam2 light chain
Increased degradation of the A181P and G308R Myo1 mutants compared to the T140I mutant or wild type Myo1 in immunoblotting assays (Fig. 4.1) suggested that their protein stability was decreased, possibly due to misfolding.
To directly test whether FSGS-associated mutations affect myosin folding and stability, we examined whether the stability of Myo1 motor mutants correlated with their ability to associate with a myosin chaperone Rng3. Rng3 is a member of the UCS family of molecular chaperones that is necessary for the motor activity of fission yeast myosin II Myo2 (Lord and Pollard, 2004; Stark et al.,
2013). Previous work showed that while Rng3 is not required for Myo1 folding, a destabilizing G308R mutation in the Myo1 motor domain promotes recruitment of
Rng3 to Myo1, which was detected using Myo1-specific light chain Cam2 tagged with mCherry (Stark et al., 2013). In myo1(G308R) cells, Rng3 and Cam2 co- localized in patch-like structures that, we suspect, represented cross-sections of eisosomes or protein aggregates.
Initially, we followed the approach taken by Stark et al. (2013) and tested whether Myo1-specific light chain Cam2 tagged with mCherry can be used to track localization of FSGS-associated Myo1 mutants in cells. We combined
Cam2-mCherry with mGFP-tagged wild type or mutant Myo1 by genetic crosses
(Fig. 4.6A-C’’, Fig. S4.4A-C’’) and examined their localization in live fission yeast
162 Chapter 4 Myo1e mutants in yeast cells. Fluorescence imaging showed that wild-type Myo1 and Myo1(G483D) co- localized with Cam2 (Fig. 4.6A-A’’, Fig. S4.4A-A’’) in 100% and 91% of endocytic actin patches, respectively (Fig, S4.5). Cam2 binds the second IQ motif of Myo1
(Sammons et al., 2011). When Cam2 binding to Myo1 was disrupted by a point mutation (Sammons et al., 2011) or by the deletion of the second IQ motif (Fig.
S4.4C-C’’), Myo1 continued to localize to actin patches, while Cam2 re-localized to mobile and stationary puncta that were clearly distinct from actin patches
(white arrows in Fig. S4C-C’’). The residual 5-12.5% co-localization between mGFP-Myo1(ΔIQ2) and Cam2-mCherry (Fig. S4.5) represented occasional fortuitous juxtaposition of actin patches and Cam2 puncta. Cam2 maintained association with Myo1 motor domain mutants and Cam2-mCherry showed robust co-localization with mGFP-tagged T140I, A181P, and G308R Myo1 mutants
(yellow arrows in Fig. 4.6B-C’’, Fig. S4.4B-B’’) in 100% of eisosomes (Fig. S4.5).
For each mutant, the amount of Cam2 in eisosomes correlated with the amount of Myo1 in eisosomes and the level of full-length Myo1 detected by immunoblotting (Fig. 4.1). In the stable T140I mutant, virtually all Cam2 co- localized with Myo1 in eisosomes, whereas in the unstable A181P and G308R mutants Cam2 localized to both eisosomes and cytoplasmic puncta (Fig. S4.5, yellow arrows in Fig. 4.6C-C’’ and Fig. S4.4B-B’’) similar to those observed in the cells expressing Myo1 mutant lacking the second IQ motif (Fig. S4.5, white arrows in Fig. S4.4C-C’’). Co-localization of Cam2 with Myo1 mutants in
163 Chapter 4 Myo1e mutants in yeast eisosomes, structures with distinctive linear organization, allowed us to use
Cam2 as a proxy for Myo1 localization.
4.5.6 Chaperone Rng3 is recruited to the A181P but not the T140I Myo1 mutant
To test if Myo1 motor domain mutants recruit Rng3, we constructed haploid yeast strains combining triple GFP-tagged Rng3 (Rng3-3xGFP) with
Cam2-mCherry and the untagged versions of either the wild type, or mutant
Myo1 (Fig. 4.6D-F’’ and Fig. S4.4D-F’’). In the wild type and myo1(G483D) cells,
Rng3-3xGFP localized to actin contractile rings (green arrows in Fig. 4.6D-D’’ and Fig. S4.4D-D’’) and cytoplasmic puncta (green arrowheads in Fig. 4.6D-D’’ and Fig. S4.4D-D’’) and did not co-localize with Cam2-mCherry in actin patches
(red arrowheads in Fig. 4.6D-D’’ and Fig. S4.4D-D’’). In contrast to the wild type cells, in myo1(A181P) and myo1(G308R) mutants, Rng3-3xGFP partially colocalized with Cam2-labeled eisosomes (yellow arrows in Fig. 4.6F-F’’ and Fig.
S4.4E-E’’), indicating recruitment of Rng3 to mutated Myo1. In addition to eisosomes, we also observed that in myo1(T140I), myo1(A181P) and myo1(G308R) cells, Cam2 and Rng3 co-localized in bright cortical puncta of unknown nature (yellow arrowheads in Fig. 4.6E-F’’ and Fig. S4.4E-E’’), which may represent fragmented eisosomes or protein aggregates. In stark contrast to the myo1(A181P) and myo1(G308R) mutants, in myo1(T140I) mutant cells Rng3 was not recruited to the eisosomes (red arrows in Fig. 4.6E-E’’), even though
Cam2 in this mutant accumulated in eisosomes to a greater degree than in
A181P or G308R mutants. We also found that in the myo1(2xIQ1IQ2) mutant
164 Chapter 4 Myo1e mutants in yeast containing normal Myo1 motor domain but duplicated IQ motifs in the neck,
Cam2 localized to eisosomes (red arrows in Fig. S4.4F’-F’’) and did not co- localize with Rng3. Thus, eisosome localization of Myo1-bound Cam2 is not always accompanied by the Rng3 recruitment.
While the differences in association with Rng3 among Myo1 motor domain mutants were readily apparent, the detailed analysis of co-localization was complicated by the variability in Cam2-mCherry localization to eisosomes in
A181P and G308R mutants combined with Rng3-3xGFP. The fluorescence signal was weak, and some cells did not exhibit clear Cam2 enrichment in eisosomes, which could be due to the effects of combining bulky mCherry and triple GFP tags. Therefore, to directly confirm association of Rng3 with A181P and G308R but not T140I Myo1 mutants in eisosomes, we employed two additional complementary approaches. We combined wild type and mutant Myo1 labeled with a fluorescent protein tag directly on the Myo1 heavy chain with Rng3 labeled with a contrasting fluorescent protein. Specifically, we combined mCherry-tagged Myo1 variants with Rng3-3xGFP and mGFP-tagged Myo1 variants with Rng3-mCherry and optimized imaging conditions to allow detection of faint mCherry-tagged proteins in live cells.
Fluorescence imaging revealed unequivocal association of Rng3 with
A181P and G308R Myo1 mutants but not T140I Myo1 mutant. In cells combining mCherry-tagged Myo1 constructs with 3xGFP-tagged Rng3 (Fig. 4.7A-C’’, Fig.
S4.6A-B’’), wild type Myo1 and Myo1(G483D) localized to patches that lacked
165 Chapter 4 Myo1e mutants in yeast any Rng3 (Fig. 4.7A-A’’ and Fig. S4.6A-A’’), while Myo1(T140I), Myo1(A181P), and Myo1(G308R) localized to eisosomes in all cells. mCherry-tagged A181P and G308R mutants in eisosomes showed robust co-localization with Rng3-
3xGFP in every cell in the population (yellow arrows in Fig. 4.7C-C’’ and Fig.
S4.6B-B’’). In contrast, despite stronger accumulation in eisosomes, Myo1(T140I) showed little co-localization with Rng3 (Fig. 4.7B-B’’), except a few rare examples of weak association of Rng3 with Myo1(T140) in only 1-2 eisosomes in less than
10% of the cells. We also noted occasional examples of Rng3 localization with
T140I, A181P, and G308R Myo1 mutants in cytoplasmic puncta, which we suspect are protein aggregates. Similar localization patterns were observed in a reciprocal experiment using mGFP-tagged Myo1 variants combined with mCherry-tagged Rng3 (Fig. 4.7D-F’’, Fig. S4.6C-E’’). Even though the Rng3- mCherry signal was extremely faint, Rng3-mCherry showed robust co- localization in eisosomes with mGFP-tagged A181P and G308R Myo1 mutants in every cell (yellow arrows in Fig. 4.7F-F’’ and Fig S4.6D-D’’). In contrast, little or no Rng3-mCherry co-localized with mGFP-tagged T140I mutant (Fig. 4.7E-E’’) or mutant with duplicated IQ motif in eisosomes (Fig. S4.6E-E’’). Rng3-mCherry also failed to co-localize with wild type and G483D mutant Myo1 in actin patches
(Fig. 4.7D-D’’ and Fig. S4.6C-C’’). Thus, association of Rng3 with Myo1 is only observed with unstable Myo1 motor domain mutants.
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4.6 Discussion
4.6.1 FSGS-associated mutations disrupt myosin-1 function
We have examined the functional effects of FSGS-associated mutations in
Myo1e by generating equivalent mutations in the fission yeast Myo1. The introduction of the T140I or A181P mutations in the S. pombe Myo1 motor domain led to defects in yeast growth under the high salt and high temperature conditions, resulting in the phenotype similar to that of the Δmyo1 strain.
Furthermore, actin patches in the Myo1 motor domain mutant strains exhibited slower patch assembly and defective patch internalization, similar to the actin patches in the Δmyo1 cells. These observations indicate that both mutations result in the loss of myosin functional activity. Taken together with our previous findings that Myo1e-null mice develop FSGS-like disease, and that cultured podocytes expressing Myo1e(A159P) exhibit defects in formation of cell-cell contacts (Bi et al., 2013; Krendel et al., 2009), our data strongly indicate that the loss of function of Myo1e is the cause of kidney disease in patients homozygous for the T119I or A159P mutations (Al-Hamed et al., 2013; Mele et al., 2011;
Sanna-Cherchi et al., 2011). Thus, our findings confirm that both mutations in
Myo1e are likely to be pathogenic, and provide evidence that a functional Myo1e motor domain is required to maintain normal glomerular filtration.
4.6.2 Molecular basis for the effects of motor domain mutations on myosin activity and stability
The two FSGS mutations examined in this study affect highly conserved
167 Chapter 4 Myo1e mutants in yeast residues in the myosin motor domain. The conserved Alanine at position 181 is located adjacent to the Switch I region, which is involved in coordinating actin binding with the conformational changes that occur during ATP hydrolysis
(Sweeney and Houdusse, 2010). We have observed that the A181P mutant is both functionally defective and unstable, exhibiting increased degradation and association with the Rng3 chaperone. A rigid Proline residue replacing the
Alanine in this position may interfere not only with the conformational changes required for motor activity but also with protein folding, thereby destabilizing the protein.
Threonine 140 is part of the P-loop consensus sequence (GXXXXGKT/S)
(Smith and Rayment, 1996), also known as a Walker A motif (Walker et al.,
1982), a conserved nucleotide-binding motif present in a variety of nucleotide- hydrolyzing enzymes (Koonin, 1993). Hydroxyl group of this amino acid residue forms a part of the cation-binding pocket that stabilizes the Mg2+ ion associated with the bound nucleotide, and, therefore, it is clearly important for ATP binding and/or hydrolysis. Indeed, mutations of this Threonine in the Walker A motif have been shown to disrupt functions of several nucleotide-hydrolyzing enzymes
(Hishida et al., 1999; Shen et al., 1994; Siddique and Figurski, 2012).
Characterization of the T93I mutation in the P-loop of kinesin provides insights into the effect of this mutation on a molecular motor protein. This mutation results in strong (rigor) binding of kinesin to microtubules (Nakata and
Hirokawa, 1995). In the kinesin mechanochemical cycle, the ATP-bound state
168 Chapter 4 Myo1e mutants in yeast corresponds to the strong microtubule-binding state, and mutations disrupting
ATP hydrolysis would result in rigor binding. Thus, the T-to-I mutation in the P- loop of kinesin likely inhibits ATP hydrolysis. Unlike kinesins, ATP-bound myosins exhibit weak binding to actin filaments (Hackney, 1996). Therefore, a mutation disrupting myosin ATP hydrolysis would be expected to result in weak binding of myosin to actin, which correlates with the observed lack of localization of Myo1-T140I to actin patches. Unlike A181P, T140I mutation has no effect on
Myo1 stability in yeast. Because T140I mutation disrupts motor domain without decreasing protein stability, this mutant is particularly interesting as it illuminates the role of myosin-1 motor domain in myosin-1 localization and function.
4.6.3 Motor domain is necessary for myosin-1 localization
Neither the T140I, nor the A181P mutant of Myo1 localized to actin patches in fission yeast, indicating that the functional motor domain is required for Myo1 localization to actin patches. In mammalian cells, wild type Myo1e is found both in the cytoplasm and on the plasma membrane, with some enrichment at the cell-cell junctions and on endocytic vesicles (Bi et al., 2013;
Krendel et al., 2007). Similar to the yeast Myo1 mutants that no longer localize to actin patches, Myo1e-A159P and Myo1e-T119I mutants fail to localize to cell-cell junctions and intracellular vesicles ((Bi et al., 2013) and our unpublished observations) and exhibit a mostly cytosolic localization. On the other hand, a
Myo1e construct completely lacking the motor and neck domains (Myo1e-tail) is able to localize to the plasma membrane and endocytic vesicles (Krendel et al.,
169 Chapter 4 Myo1e mutants in yeast
2007). However, Myo1e-tail does not completely recapitulate the normal localization of the full-length Myo1e, showing an exaggerated enrichment in the plasma membrane compared to the full-length Myo1e (Bi et al., 2013). Overall, a functional motor domain appears to be necessary for the normal intracellular localization of both human Myo1e and yeast Myo1.
In addition to the lack of co-localization between the disease-associated
Myo1 mutants and actin patches, we observed an unexpected enrichment of the mutant Myo1 variants in eisosomes. We hypothesize that when mutations in the
Myo1 motor domain disrupt its ability to localize to actin patches, interactions of the Myo1 tail with the lipids, such as PIP2 (Kabeche et al., 2014), or proteins that are enriched in eisosomes result in targeting of Myo1 to eisosomes. The connection between the lack of motor activity and eisosomal localization of Myo1 is further substantiated by the observation that a previously characterized severe motor domain mutant of Myo1, G308R, also localized to eisosomes while the mild, patch-localizing motor domain mutant G483D did not localize to eisosomes.
The observed role for motor activity in determining intracellular localization of
Myo1 is in agreement with the finding that Myo1 lacking motor domain also localizes to eisosomes (R.T.C., manuscript in preparation).
4.6.4 Myo1e motor domain functions in the kidney
The fact that both FSGS-associated mutations affect motor domain function underscores the importance of Myo1e motor activity for its roles in renal filtration. The understanding of how Myo1e motor activity contributes to the
170 Chapter 4 Myo1e mutants in yeast normal structure and function of the glomerular filtration barrier is still incomplete.
Some of the defects caused by the motor domain mutations could be due to the mis-localization of the myosin molecule. However, it is unlikely that the motor domain function is limited to determining Myo1e localization. Indeed, a headless
Myo1e construct inhibits endocytosis in a dominant-negative fashion, indicating that it is unable to functionally replace full-length Myo1e (Krendel et al., 2007).
Class I myosins may act as multifunctional linkers between the actin filaments that interact with the motor domain and the plasma membrane and membrane-associated protein complexes that bind to the tail domain (Arif et al.,
2011; Bi et al., 2013; McConnell and Tyska, 2010). Myosin motor domain activity and the ability to bind to actin are likely to be important for the processes that require coordination of membrane deformation with actin filament recruitment, including cell shape changes, endocytosis, assembly of cell adhesion complexes, and regulation of membrane tension. Indeed, studies in cultured cells have demonstrated contributions of Myo1e to endocytosis, regulation of membrane tension, and cell adhesion dynamics (Cheng et al., 2012; Gupta et al., 2013;
Krendel et al., 2009; Nambiar et al., 2009). Furthermore, podocytes that express
Myo1e(A159P) as the sole source of Myo1e exhibited defects in actin assembly at cell-cell junctions (Bi et al., 2013). In the absence of a functional motor domain,
Myo1e may be unable to link the plasma membrane to the underlying actin cytoskeleton or recruit actin filaments to specific membrane subdomains, leading to defects in podocyte adhesion, endocytosis, and cell shape regulation.
171 Chapter 4 Myo1e mutants in yeast
Disruption of these processes leads to defects in glomerular filtration and FSGS pathogenesis (Faul et al., 2007; George and Holzman, 2012; Lennon et al., 2014;
Soda and Ishibe, 2013).
In conclusion, we have established fission yeast S. pombe as an effective model system for testing the effects of kidney disease-associated mutations on myosin-1 function. By using this simple but powerful system, in which localization and function of myosin-1 are well defined, we have gained insights into the role of myosin-1 motor activity that are also applicable to human kidney biology.
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4.7 References
Al-Hamed, M. H., Al-Sabban, E., Al-Mojalli, H., Al-Harbi, N., Faqeih, E., Al Shaya, H., Alhasan, K., Al-Hissi, S., Rajab, M., Edwards, N. et al. (2013). A molecular genetic analysis of childhood nephrotic syndrome in a cohort of Saudi Arabian families. J Hum Genet 58, 480-9. Arif, E., Wagner, M. C., Johnstone, D. B., Wong, H. N., George, B., Pruthi, P. A., Lazzara, M. J. and Nihalani, D. (2011). Motor protein Myo1c is a podocyte protein that facilitates the transport of slit diaphragm protein Neph1 to the podocyte membrane. Mol Cell Biol 31, 2134- 50. Bahler, J., Wu, J. Q., Longtine, M. S., Shah, N. G., McKenzie, A., 3rd, Steever, A. B., Wach, A., Philippsen, P. and Pringle, J. R. (1998). Heterologous modules for efficient and versatile PCR-based gene targeting in Schizosaccharomyces pombe. Yeast 14, 943-51. Balasubramanian, M. K., McCollum, D., Chang, L., Wong, K. C., Naqvi, N. I., He, X., Sazer, S. and Gould, K. L. (1998). Isolation and characterization of new fission yeast cytokinesis mutants. Genetics 149, 1265-75. Bement, W. M., Wirth, J. A. and Mooseker, M. S. (1994). Cloning and mRNA expression of human unconventional myosin-IC. A homologue of amoeboid myosins-I with a single IQ motif and an SH3 domain. J Mol Biol 243, 356-63. Berg, J. S., Powell, B. C. and Cheney, R. E. (2001). A millennial myosin census. Mol Biol Cell 12, 780-94. Bi, J., Chase, S. E., Pellenz, C. D., Kurihara, H., Fanning, A. S. and Krendel, M. (2013). Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am J Physiol Renal Physiol 305, F532-44. Chase, S. E., Encina, C. V., Stolzenburg, L. R., Tatum, A. H., Holzman, L. B. and Krendel, M. (2012). Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am J Physiol Renal Physiol 303, F1099-106. Cheng, J., Grassart, A. and Drubin, D. G. (2012). Myosin 1E coordinates actin assembly and cargo trafficking during clathrin-mediated endocytosis. Mol Biol Cell 23, 2891-904. El Mezgueldi, M., Tang, N., Rosenfeld, S. S. and Ostap, E. M. (2002). The kinetic mechanism of Myo1e (human myosin-IC). J Biol Chem 277, 21514-21. Faul, C., Asanuma, K., Yanagida-Asanuma, E., Kim, K. and Mundel, P. (2007). Actin up: regulation of podocyte structure and function by components of the actin cytoskeleton. Trends Cell Biol 17, 428-37. Feeser, E. A., Ignacio, C. M., Krendel, M. and Ostap, E. M. (2010). Myo1e binds anionic phospholipids with high affinity. Biochemistry 49, 9353-60. George, B. and Holzman, L. B. (2012). Signaling from the podocyte intercellular junction to the actin cytoskeleton. Semin Nephrol 32, 307-18. Greka, A. and Mundel, P. (2012). Cell biology and pathology of podocytes. Annu Rev Physiol 74, 299-323. Gupta, P., Gauthier, N. C., Cheng-Han, Y., Zuanning, Y., Pontes, B., Ohmstede, M., Martin, R., Knolker, H. J., Dobereiner, H. G., Krendel, M. et al. (2013). Myosin 1E localizes to actin polymerization sites in lamellipodia, affecting actin dynamics and adhesion formation. Biol Open 2, 1288-99. Hackney, D. D. (1996). The kinetic cycles of myosin, kinesin, and dynein. Annu Rev Physiol 58, 731-50. Hishida, T., Iwasaki, H., Yagi, T. and Shinagawa, H. (1999). Role of walker motif A of RuvB protein in promoting branch migration of holliday junctions. Walker motif a mutations affect Atp binding, Atp hydrolyzing, and DNA binding activities of Ruvb. J Biol Chem 274, 25335-42. Kabeche, R., Baldissard, S., Hammond, J., Howard, L. and Moseley, J. B. (2011). The filament-forming protein Pil1 assembles linear eisosomes in fission yeast. Mol Biol Cell 22, 4059-67.
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Kabeche, R., Roguev, A., Krogan, N. J. and Moseley, J. B. (2014). A Pil1-Sle1-Syj1- Tax4 functional pathway links eisosomes with PI(4,5)P2 regulation. J Cell Sci 127, 1318-26. Koonin, E. V. (1993). A superfamily of ATPases with diverse functions containing either classical or deviant ATP-binding motif. J Mol Biol 229, 1165-74. Krendel, M., Kim, S. V., Willinger, T., Wang, T., Kashgarian, M., Flavell, R. A. and Mooseker, M. S. (2009). Disruption of Myosin 1e promotes podocyte injury. J Am Soc Nephrol 20, 86-94. Krendel, M., Osterweil, E. K. and Mooseker, M. S. (2007). Myosin 1E interacts with synaptojanin-1 and dynamin and is involved in endocytosis. FEBS Lett 581, 644-50. Lee, W. L., Bezanilla, M. and Pollard, T. D. (2000). Fission yeast myosin-I, Myo1p, stimulates actin assembly by Arp2/3 complex and shares functions with WASp. J Cell Biol 151, 789-800. Lennon, R., Randles, M. J. and Humphries, M. J. (2014). The importance of podocyte adhesion for a healthy glomerulus. Front Endocrinol (Lausanne) 5, 160. Lord, M. and Pollard, T. D. (2004). UCS protein Rng3p activates actin filament gliding by fission yeast myosin-II. J Cell Biol 167, 315-25. McConnell, R. E. and Tyska, M. J. (2010). Leveraging the membrane - cytoskeleton interface with myosin-1. Trends Cell Biol 20, 418-26. Mele, C., Iatropoulos, P., Donadelli, R., Calabria, A., Maranta, R., Cassis, P., Buelli, S., Tomasoni, S., Piras, R., Krendel, M. et al. (2011). MYO1E mutations and childhood familial focal segmental glomerulosclerosis. N Engl J Med 365, 295-306. Moore, J. R., Leinwand, L. and Warshaw, D. M. (2012). Understanding cardiomyopathy phenotypes based on the functional impact of mutations in the myosin motor. Circ Res 111, 375- 85. Moreira, K. E., Walther, T. C., Aguilar, P. S. and Walter, P. (2009). Pil1 controls eisosome biogenesis. Mol Biol Cell 20, 809-18. Munn, A. L., Stevenson, B. J., Geli, M. I. and Riezman, H. (1995). end5, end6, and end7: mutations that cause actin delocalization and block the internalization step of endocytosis in Saccharomyces cerevisiae. Mol Biol Cell 6, 1721-42. Nakata, T. and Hirokawa, N. (1995). Point mutation of adenosine triphosphate-binding motif generated rigor kinesin that selectively blocks anterograde lysosome membrane transport. J Cell Biol 131, 1039-53. Nambiar, R., McConnell, R. E. and Tyska, M. J. (2009). Control of cell membrane tension by myosin-I. Proc Natl Acad Sci U S A 106, 11972-7. Olivera-Couto, A., Grana, M., Harispe, L. and Aguilar, P. S. (2011). The eisosome core is composed of BAR domain proteins. Mol Biol Cell 22, 2360-72. Ouderkirk, J. L. and Krendel, M. (2014). Non-muscle myosins in tumor progression, cancer cell invasion, and metastasis. Cytoskeleton (Hoboken) 71, 447-63. Redowicz, M. J. (2002). Myosins and pathology: genetics and biology. Acta Biochim Pol 49, 789-804. Roseaulin, L. C., Noguchi, C., Martinez, E., Ziegler, M. A., Toda, T. and Noguchi, E. (2013). Coordinated degradation of replisome components ensures genome stability upon replication stress in the absence of the replication fork protection complex. PLoS Genet 9, e1003213. Sammons, M. R., James, M. L., Clayton, J. E., Sladewski, T. E., Sirotkin, V. and Lord, M. (2011). A calmodulin-related light chain from fission yeast that functions with myosin-I and PI 4-kinase. J Cell Sci 124, 2466-77. Sanna-Cherchi, S., Burgess, K. E., Nees, S. N., Caridi, G., Weng, P. L., Dagnino, M., Bodria, M., Carrea, A., Allegretta, M. A., Kim, H. R. et al. (2011). Exome sequencing identified MYO1E and NEIL1 as candidate genes for human autosomal recessive steroid-resistant nephrotic syndrome. Kidney Int 80, 389-96. Shen, H., Yao, B. Y. and Mueller, D. M. (1994). Primary structural constraints of P-loop of mitochondrial F1-ATPase from yeast. J Biol Chem 269, 9424-8.
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Siddique, A. and Figurski, D. H. (2012). Different phenotypes of Walker-like A box mutants of ParA homolog IncC of broad-host-range IncP plasmids. Plasmid 68, 93-104. Sirotkin, V., Beltzner, C. C., Marchand, J. B. and Pollard, T. D. (2005). Interactions of WASp, myosin-I, and verprolin with Arp2/3 complex during actin patch assembly in fission yeast. J Cell Biol 170, 637-48. Sirotkin, V., Berro, J., Macmillan, K., Zhao, L. and Pollard, T. D. (2010). Quantitative analysis of the mechanism of endocytic actin patch assembly and disassembly in fission yeast. Mol Biol Cell 21, 2894-904. Smith, C. A. and Rayment, I. (1996). Active site comparisons highlight structural similarities between myosin and other P-loop proteins. Biophys J 70, 1590-602. Snaith, H. A., Samejima, I. and Sawin, K. E. (2005). Multistep and multimode cortical anchoring of tea1p at cell tips in fission yeast. EMBO J 24, 3690-9. Soda, K. and Ishibe, S. (2013). The function of endocytosis in podocytes. Curr Opin Nephrol Hypertens 22, 432-8. Stark, B. C., James, M. L., Pollard, L. W., Sirotkin, V. and Lord, M. (2013). UCS protein Rng3p is essential for myosin-II motor activity during cytokinesis in fission yeast. PLoS One 8, e79593. Sweeney, H. L. and Houdusse, A. (2010). Structural and functional insights into the Myosin motor mechanism. Annu Rev Biophys 39, 539-57. Tajsharghi, H. and Oldfors, A. (2013). Myosinopathies: pathology and mechanisms. Acta Neuropathol 125, 3-18. Walker, J. E., Saraste, M., Runswick, M. J. and Gay, N. J. (1982). Distantly related sequences in the alpha- and beta-subunits of ATP synthase, myosin, kinases and other ATP- requiring enzymes and a common nucleotide binding fold. EMBO J 1, 945-51. Walther, T. C., Aguilar, P. S., Frohlich, F., Chu, F., Moreira, K., Burlingame, A. L. and Walter, P. (2007). Pkh-kinases control eisosome assembly and organization. EMBO J 26, 4946-55. Whitacre, J., Davis, D., Toenjes, K., Brower, S. and Adams, A. (2001). Generation of an isogenic collection of yeast actin mutants and identification of three interrelated phenotypes. Genetics 157, 533-43. Wong, K. C., Naqvi, N. I., Iino, Y., Yamamoto, M. and Balasubramanian, M. K. (2000). Fission yeast Rng3p: an UCS-domain protein that mediates myosin II assembly during cytokinesis. J Cell Sci 113 ( Pt 13), 2421-32. Wu, J. Q., Kuhn, J. R., Kovar, D. R. and Pollard, T. D. (2003). Spatial and temporal pathway for assembly and constriction of the contractile ring in fission yeast cytokinesis. Dev Cell 5, 723-34. Wu, J. Q. and Pollard, T. D. (2005). Counting cytokinesis proteins globally and locally in fission yeast. Science 310, 310-4.
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Figure 4.1. Kidney disease-associated mutations in S. pombe Myo1 result in growth defects and decreased protein stability.
176 Chapter 4 Myo1e mutants in yeast
Figure 4.1. Kidney disease-associated mutations in S. pombe Myo1 result in growth defects and decreased protein stability. (A) Domain maps of H. sapiens Myo1e and S. pombe Myo1. IQ – light chain-binding IQ motif, TH – Tail
Homology domain, SH3 – Src Homology 3 domain, CA – Central-Acidic domain.
(B) Alignment of amino acid sequences of the N-terminal portions of H. sapiens
Myosin 1e (Myo1e) and S. pombe Myosin 1 (Myo1). Red bold font indicates the sites of the two FSGS-associated mutations. The conserved P-loop motif and
Switch I motifs are underlined. The T119I and A159P mutations in H.s. Myo1e are equivalent to the T140I and A181P mutations in S.p. Myo1, respectively. (C)
Analysis of the salt and temperature sensitivity of the wild type and untagged myo1 mutant strains. The myo1(T140I), myo1(A181P) and Δmyo1 strains fail to grow in the presence of high salt at high temperature (YE5S+1M KCl at 36°C) and grow but stain darkly with Phloxine B (PB) on minimal EMM medium at 25° and 36°C, indicating decreased cell viability. (D-G) Western blot analysis of yeast extracts using anti-GFP antibody to determine protein stability of the mGFP- tagged Myo1 constructs. (D) Similar amounts of the full-length Myo1-mGFP (FL, arrow) are detected in cells expressing mGFP-tagged wild type, T140I or G483D mutant Myo1 while the amount of the full length myosin is reduced in the strains expressing the A181P and G308R Myo1 mutants. Arrowhead indicates the ~40 kDa degradation product. An asterisk indicates a lower ~30 kDa molecular weight degradation product that is enriched in the A181P and G308R mutants. No signal is detected in Δmyo1 strain. Coomassie-stained gel (bottom panel) served as the
177 Chapter 4 Myo1e mutants in yeast loading control. (E, F) Quantification of (E) total band intensity and (F) full-length band intensity normalized to the wild type. N=5 blots from 3 independent experiments. Error bars represent S.D. (G) The background-subtracted whole cell intensity of mGFP-tagged Myo1 variants in cells before lysis. N= 5 cells. Error bars represent S.D.
178 Chapter 4 Myo1e mutants in yeast
Figure 4.2. Motor domain mutations disrupt Myo1 localization to actin patches.
179 Chapter 4 Myo1e mutants in yeast
Figure 4.2. Motor domain mutations disrupt Myo1 localization to actin patches. Confocal images show the localization of (left panels, A-F) mGFP- tagged (A) wild-type or (B-F) mutant Myo1, (middle panels, A’-F’) mCherry- tagged Fimbrin (Fim1-mCherry), and (right panels, A’’-F’’) merge of Myo1-mGFP
(green) and Fim1-mCherry (red) images. Wild type Myo1 and Myo1(G483D) co- localize with Fim1-mCherry in actin patches (yellow arrowheads), while
Myo1(T140I), Myo1(A181P), and Myo1(G308R) localize to eisosomes (green arrows) and are absent from actin patches (red arrowheads). The images represent maximum intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4 µm intervals. Scale bar, 10 µm.
180 Chapter 4 Myo1e mutants in yeast
Figure 4.3. Motor domain mutations disrupt dynamics of endocytic actin patches visualized with mCherry-tagged fimbrin Fim1.
181 Chapter 4 Myo1e mutants in yeast
Figure 4.3. Motor domain mutations disrupt dynamics of endocytic actin patches visualized with mCherry-tagged fimbrin Fim1. (A–F) Time courses of
(upper panels) average fluorescence intensity of (green, solid circles) wild type and mutant Myo1-mGFP and (red, open squares) Fim1-mCherry in actin patches and (lower panels) raw (thin lines) and average (thick lines) distances traveled by
Fim1-mCherry or Myo1-mGFP in patches. (G-I) Bar graphs showing (G) the total lifetime, assembly time and disassembly time, (H) percentage of internalization and (I) mean peak intensities of Fim1-mCherry patches in wild type and mutant myo1 strains. Note increased total lifetime, assembly and disassembly times, and decreased internalization of Fim1-mCherry patches in the T140I, A181P, G308R myo1 mutant, and Δmyo1 strains. In all panels, error bars represent S.D. (A-G and I), N= 5-12 patches. (H), N=23-43 patches in 3-6 cells. The asterisks indicate statistical significance: *, p < 0.05; ** p < 0.01.
182 Chapter 4 Myo1e mutants in yeast
Figure 4.4. Myo1 motor domain mutants co-localize with the eisosome marker Fhn1.
183 Chapter 4 Myo1e mutants in yeast
Figure 4.4. Myo1 motor domain mutants co-localize with the eisosome marker Fhn1. Confocal images show the localization of (left panels, A-E) mGFP- tagged (A) wild-type or (B-E) mutant Myo1, (middle panels, A’-E’) mCherry- tagged Fhn1 (Fhn1-mCherry), and (right panels, A’’-E’’) merge of Myo1-mGFP and Fhn1-mCherry images. The wild type and Myo1(G483D) localize to actin patches (green arrowheads) that are distinct from Fhn1-mCherry in eisosomes
(red arrows). The T140I, A181P, and G308R Myo1 mutants co-localize with Fhn1 in eisosomes (yellow arrows). The images represent maximum intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4 µm intervals. Scale bar, 10 µm.
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Figure 4.5. Myo1 mutants do not co-localize with filamentous actin labeled by Lifeact-mCherry.
185 Chapter 4 Myo1e mutants in yeast
Figure 4.5. Myo1 mutants do not co-localize with filamentous actin labeled by Lifeact-mCherry. Confocal images show the localization of (left panels, A-D) mGFP-tagged (A) wild type and (B-D) mutant Myo1, (panels A’-D’) Lifeact- mCherry, (panels A’’-D’’) merge of Myo1-mGFP and Lifeact-mCherry images, and (right panels, A’’’-D’’’) line scans in green (green lines) and red (red lines) channels along the white lines indicated on the merged images in panels A’’-D’’.
Wild type Myo1 co-localizes with Lifeact in actin patches (yellow arrowheads) but not in actin cables (red arrows). The T140I and A181P Myo1 mutants localize to eisosomes (green arrows, E in line scans) that are distinct from actin cables (red arrows, C in line scans), patches (open red arrowheads, P in line scans) and rings (filled red arrowheads in B’-B’’). The images represent maximum intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4 µm intervals. Scale bar, 10 µm.
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Figure 4.6. Myo1 light chain Cam2 co-localizes with Myo1 mutants and co- localizes with Rng3 in cells expressing A181P but not the T140I Myo1 mutants.
187 Chapter 4 Myo1e mutants in yeast
Figure 4.6. Myo1 light chain Cam2 co-localizes with Myo1 mutants and co- localizes with Rng3 in cells expressing A181P but not the T140I Myo1 mutants. (A-C’’) Confocal images show the localization of (left panels, A-C) mGFP-tagged (A), wild-type and (B-C), mutant Myo1, (middle panels, A’-C’) mCherry-tagged Cam2 (Cam2-mCherry), and (right panels, A’’-C’’) merge of
Myo1-mGFP and Cam2-mCherry images. Cam2-mCherry co-localizes with mGFP-tagged wild-type Myo1 in patches (yellow arrowheads) and with mGFP- tagged T140I and A181P Myo1 mutants in eisosomes (yellow arrows). In A181P mutant, Cam2 also localizes to cytoplasmic puncta that are devoid of Myo1
(white arrows). (D-F’’) Confocal images show the localization of (left panels, D-F) triple GFP-tagged Rng3 (Rng3-3xGFP), (middle panels, D’-F’) mCherry-tagged
Cam2 (Cam2-mCherry), and (right panels D’’-F’’) merge of Rng3-3xGFP and
Cam2-mCherry images in yeast strains expressing untagged wild type or kidney disease-associated Myo1 variants. In the wild type cells, Rng3 localizes to the contractile rings (green arrows) and cytoplasmic spots of unknown nature (green arrowheads) that are distinct from Cam2 in actin patches (red arrowheads). Rng3 does not co-localize with Cam2 in eisosomes (red arrows in E’-E’’) in the T140I mutant, but co-localizes with Cam2 in eisosomes (yellow arrows in F-F’’) in the
A181P mutant. Cam2 and Rng3 also co-localize in cortical puncta in A181P and, occasionally, T140I mutant (yellow arrowheads in E-F’’). The images represent maximum intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4 µm intervals. Scale bar, 10 µm.
188 Chapter 4 Myo1e mutants in yeast
Figure 4.7. Chaperone Rng3 is recruited to the A181P but not the T140I
Myo1 mutant in eisosomes.
189 Chapter 4 Myo1e mutants in yeast
Figure 4.7. Chaperone Rng3 is recruited to the A181P but not the T140I
Myo1 mutant in eisosomes. (A-C’’) Confocal images show the localization of
(left panels, A-C) triple GFP-tagged Rng3 (Rng3-3xGFP), (middle panels, A’-C’) mCherry-tagged Myo1 variants (Myo1-mCherry), and (right panels A’’-C’’) merge of Rng3-3xGFP and Myo1-mCherry images.
Wild type Myo1 in patches (red arrowheads) and Myo1(T140I) in eisosomes (red arrows) are distinct from Rng3 puncta (green arrowheads). Rng3 co-localizes with A181P Myo1 mutant in eisosomes (yellow arrows). (D-F’’) Confocal images show the localization of (left panels, D-F) mGFP-tagged Myo1 variants (Myo1- mGFP), (middle panels, D’-F’) mCherry-tagged Rng3 (Rng3-mCherry), and (right panels D’’-F’’) merge of Myo1-mGFP variants and Rng3-mCherry images. Rng3 co-localizes with Myo1(A181P) in eisosomes (yellow arrows) but not with
Myo1(T140I) in eisosomes (green arrows) or wild type Myo1 in patches (green arrowheads). The images represent maximum intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4
µm intervals. Scale bar, 10 µm.
190 Chapter 4 Myo1e mutants in yeast
Supplemental Information Table S4.1. S. pombe strains used in this study.
Strain Genotype Source Wild type and untagged myo1 mutants TP192 h- ade6-M216 leu1-32 ura4-D18 his3-D1 Δmyo1 Sirotkin et al. (2005) VS685B h- ade6-M216 leu1-32 ura4-D18 his3-D1 myo1 S. Forsburg VS1139-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 Stark et al. myo1(G483D) (2013) VS1191-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 Stark et al. myo1(G308R) (2013) VS1431 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G308R) pUR-myo1+::ura4+ VS1828-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(A181P) VS1829-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 myo1(T140I) This study mGFP-tagged Myo1 strains VS1254 h- ade6-M216 leu1-32 ura4-D18 his3-D1 Stark et al. myo1(G483D)-mGFP-kanMX6 (2013) VS1255 h- ade6-M216 leu1-32 ura4-D18 his3-D1 Stark et al. myo1(G308R)-mGFP-kanMX6 (2013) VS1256 h+ ade6-M210 leu1-32 ura4-D18 his7-366 Stark et al. myo1-mGFP-kanMX6 (2013) VS1432 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(308)-mGFP-kanMX6 pUR-myo1+::ura4+ VS1456-2 h+ ade6-M210 leu1-32 ura4-D18 his3-D1 This study myo1-mGFP-kanMX6 VS1844-2 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(T140I)-mGFP-kanMX6 VS1869-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(A181P)-mGFP-KanMX6 mCherry-tagged Myo1 strains VS1457C h+ ade6-M210 leu1-32 ura4-D18 his3-D1 This study myo1-mCherry-kanMX6 VS1843-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G483D)-mCherry-natMX6 VS1867-2 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(T140I)-mCherry-natMX6 VS1868-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(A181P)-mCherry-natMX6 VS1887-2 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G308R)-mCherry-natMX6
191 Chapter 4 Myo1e mutants in yeast
Fim1-mCherry strains VS1263-7D h+ ade6-M216 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1D::kanMX6 VS1269-2A h+ ade6-M216 leu1-32 ura4-D18 his- This study fim1-mCherry-natMX6 myo1-mGFP-kanMX6 VS1270-3B h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1(G308R)-mGFP-kanMX6 VS1271-12A h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1(G483D)-mGFP-kanMX6 VS1872-3A h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1(T140I)-mGFP-kanMX6 VS1873-7A h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1(A181P)-mGFP-kanMX6 VS1890-9D h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1(A181P)-mGFP-kanMX6 VS1892-6D h? ade6-M210 leu1-32 ura4-D18 his3-D1 This study fim1-mCherry-natMX6 myo1(A181P)-mGFP-kanMX6 Cam2-mCherry strains VS1274 h- ade6-M216 leu1-32 ura4-D18 his3-D1 Sammons et al. cam2-mCherry-kanMX6 (2011) VS1275 h+ ade6-M210 leu1-32 ura4-D18 his3-D1 Sammons et al. cam2-mCherry-kanMX6 (2011) VS1285-3A h? ade6-M216 leu1-32 ura4-D18 his3-D1 kanMX6- This study Pmyo1-mGFP-myo1DIQ2 cam2-mCherry-kanMX6 VS1289-1A h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1-mGFP-kanMX6 cam2-mCherry-kanMX6 VS1290-6A h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G483D)-mGFP-kanMX6 cam2-mCherry- kanMX6 VS1465-5C h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G308R)-mGFP-kanMX6 cam2-mCherry- kanMX6 VS1874-1B h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study cam2-mCherry-kan myo1(T140I)-mGFP-kanMX6 VS1875-2B h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study cam2-mCherry-kan myo1(A181P)-mGFP-kanMX6 Fhn1-mCherry strains VS1718 h- ade6+ ura4+ leu1+ his+ fhn1-mCherry-natMX6 J. Moseley VS1747-9B h+ ade6-M210 ura4+ leu1+ his+ fhn1-mCherry- This study natMX6 VS1747-10B h- ade6-M210 ura4+ leu1+ his+ fhn1-mCherry-natMX6 This study VS1876-4C h? ade6-M210 leu1-32 ura4+ his3-D1 This study fhn1-mCherry-Nat myo1(T140I)-mGFP-kanMX6 VS1877-3C h? ade6-M210 leu1+ ura4+ his3-D1 This study
192 Chapter 4 Myo1e mutants in yeast
fhn1-mCherry-Nat myo1(A181P)-mGFP-kanMX6 VS1878-2D h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study fhn1-mCherry-Nat myo1(483)-mGFP-kanMX6 VS1881-2B h? ade6-M210 leu1-32 ura4+ his+ This study fhn1-mCherry -Nat kanMX6-Pmyo1-mGFP-myo1 VS1883-5D h? ade6-M210 leu1+ ura4-D18 his+ This study fhn1-mCherry-Nat myo1(308)-mGFP-kanMX6
Rng3-3xGFP strains VS1216-8 h+ ade6-M210 leu1-32 ura4-D18 his3-D1 This study myo1(2xIQ1IQ2) VS1348-1 h+ ade6-M210 leu1-32 ura4-D18 his3-D1 This study rng3-3xGFP-MX6 VS1464-3A h- ade6-M216 leu1-32 Ura4-D18 his3-D1 This study cam2-mCherry-kanMX6 rng3-3xGFP-kanMX6 VS1464-6C h+ ade6-M210 leu1-32 Ura4-D18 his3-D1 This study cam2-mCherry-kanMX6 rng3-3xGFP-kanMX6 VS1893-2A h- M216 leu1-32 ura4-D18 his3-D1 This study myo1-mCherry-kanMX6 VS1894-4D h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(T140I)-mCherry-natMX6 rng3-3xGFP-kanMX6 VS1895- h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study 12D myo1(A181P)-mCherry-natMX6 rng3-3xGFP-kanMX6 VS1896-2A h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G483D)-mCherry-natMX6 rng3-3xGFP-kanMX6 VS1905- h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study 10D myo1(G308R)-mCherry-natMX6 rng3-3xGFP-kanMX6 VS1906-9D h? M216 leu1-32 ura4-D18 his3-D1 This study myo1-mCherry-kanMX6 rng3-3xGFP-kanMX6 VS1914-5D h? ade6-M216 leu1-32 ura4-D18 his3-D1 myo1(T140I) This study cam2-mCherry-kanMX6 rng3-3xGFP-kanMX6 VS1915-8D h? ade6-M210 leu1-32 ura4-D18 his3-D1 This study myo1(A181P) cam2-mCherry-kanMX6 rng3-3xGFP- kanMX6 VS1916-4A h? ade6-M210 leu1-32 ura4-D18 his3-D1 This study myo1(2xIQ1IQ2) cam2-mCherry-kanMX6 rng3- 3xGFP-kanMX6 VS1917-4B h? ade6-M216 leu1-32 ura4-D18 his3-D1 This study myo1(G483D) cam2-mCherry-kan MX6 rng3-3xGFP- kan MX6 VS1919 h+ ade6-M216 leu1-32 ura4-D18 his3-D1 Stark et al. (ML939) myo1(G308R) cam2-mCherry-kanMX6 rng3-3xGFP- (2013) kanMX6
193 Chapter 4 Myo1e mutants in yeast
Rng3-mCherry strains VS1217-1 h- ade6-M216 leu1-32 ura4-D18 his3-D1 This study kanMX6-mGFP-Pmyo1-myo1(2xIQ1IQ2) VS1920 h+ ade6-M216 leu1-32 ura4-D18 his7-366 M. Lord (ML852) myo2-E1 rng3-mCherry-kanR VS1960-1D h- ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR VS1961-6A h+ ade6-M210 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR VS1955- h? ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study 11D kanR myo1-mGFP-kanMX6 VS1956-3B h? ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR kanMX6-mGFP-Pmyo1-myo1(2xIQ1IQ2) VS1957-8D h? ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR myo1(G483D)-mGFP-kanMX6 VS1958-5D h? ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR myo1(G308R)-mGFP-kanMX6 VS1959-10B h? ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR myo1(A181P)-mGFP-kanMX6 VS1972-1D h? ade6-M216 leu1-32 ura4-D18 his- rng3-mCherry- This study kanR myo1(T140I)-mGFP-kanMX6 Lifeact-mCherry strains VS1529 h- ade6-M216 leu1-32 ura4-D18 leu1::Pact1-Lifeact- Huang et al. (MBY 6844) mCherry-leu1+ (2012) VS1669-3B h+ ade6-M210 ura4-D18 leu1-32::Pact1-Lifeact- This study mCherry-leu1+ VS1947-4D h? ade6-M210 ura4-D18 myo1(T140I)-mGFP-kanMX6 This study leu::Pact1-Lifeact-mCherry-leu+ VS1948-5C h? ade6-M210 ura4-D18 myo1(A181P)-mGFP- This study kanMX6 leu::Pact1-Lifeact-mCherry-leu+ VS1953-5A h? ade6-M216 ura4-D18 his3-D1 Myo1-mGFP- This study kanMX6 leu::Pact1-Lifeact-mCherry-leu+ VS1954-4A h? ade6-M21? ura4-D18 his3-D1 myo1Δ::kanMX6 This study leu1-32::Pact1-Lifeact-mCherry-leu+
References Huang, J., Huang, Y., Yu, H., Subramanian, D., Padmanabhan, A., Thadani, R., Tao, Y., Tang, X., Wedlich-Soldner, R., Balasubramanian, M.K. (2012). J Cell Biol. 199(5): 831-47. doi: 10.1083/jcb.201209044. Sammons, M. R., James, M. L., Clayton, J. E., Sladewski, T. E., Sirotkin, V. and Lord, M. (2011). A calmodulin-related light chain from fission yeast that functions with myosin-I and PI 4-kinase. J Cell Sci 124, 2466-77.
194 Chapter 4 Myo1e mutants in yeast
Sirotkin, V., Beltzner, C. C., Marchand, J. B. and Pollard, T. D. (2005). Interactions of WASp, myosin-I, and verprolin with Arp2/3 complex during actin patch assembly in fission yeast. J Cell Biol 170, 637-48. Stark, B. C., James, M. L., Pollard, L. W., Sirotkin, V. and Lord, M. (2013). UCS protein Rng3p is essential for myosin-II motor activity during cytokinesis in fission yeast. PLoS One 8, e79593.
195 Chapter 4 Myo1e mutants in yeast
Figure S4.1. The A181P but not T140I mutation reduces the stability of
Myo1 in cell extracts before and after cycloheximide treatment, related to
Figure 4.1.
196 Chapter 4 Myo1e mutants in yeast
Figure S4.1. The A181P but not T140I mutation reduces the stability of
Myo1 in cell extracts before and after cycloheximide treatment, related to
Figure 4.1. (A) Ratio of full length band intensity to total intensity of all bands detected by Western blotting for each mGFP-tagged Myo1 variant. (B) Western blot showing mGFP-tagged Myo1 variants detected with anti-GFP antibody in extracts of cells before and after 24-hour treatment with 0.1 mg/ml cycloheximide
(CHX). The arrow indicates full-length (FL) Myo1-mGFP. The arrowhead indicates a 40 kDa degradation product. The asterisk indicates a 30 kDa degradation product that is only present in T140I, A181P and G308R Myo1 mutants. Coomassie-stained gel (Coom.) serves as a loading control.
197 Chapter 4 Myo1e mutants in yeast
Figure S4.2. Motor domain mutations result in re-localization of Myo1 from actin patches to eisosomes, related to Figure 4.2.
198 Chapter 4 Myo1e mutants in yeast
Figure S4.2. Motor domain mutations result in re-localization of Myo1 from actin patches to eisosomes, related to Figure 4.2. (A) Analysis of co- localization between mGFP-tagged Myo1 variants and Fim1-mCherry in endocytic actin patches. Fraction of Fim1-mCherry patches that contained Myo1- mGFP at any point during patch lifetime was measured in time series in single confocal sections through the middle of the cells. Error bars represent S.D. and asterisks indicate statistical significance (p<0.001). N= 38-55 patches in 5 cells.
(B-E) Average line scans of fluorescence intensity of mGFP-tagged (blue circles)
T140I, (red squares) A181P, and (green triangles) G308R Myo1 mutants in eisosomes. An asterisk indicates statistically greater (p<10-5) peak accumulation in eisosomes for T140I mutant compared to A181P and G308R mutants. Raw line scans across 10 eisosomes for (C) T140I, (D) A181P, and (E) G308R Myo1 mutants were subtracted for extracellular background, aligned to peak intensity
(zero distance), and averaged at each point. The error bars represent S.D. The insets show examples of line scans (yellow lines) across eisosomes in single confocal sections through the top surface of the cells (outlined with white dashed lines). Scale bar, 2 μm.
199 Chapter 4 Myo1e mutants in yeast
Figure S4.3. Recruitment of mutant Myo1 to eisosomes has no effect on the number of eisosomes in cells, related to Figure 4.4.
200 Chapter 4 Myo1e mutants in yeast
Figure S4.3. Recruitment of mutant Myo1 to eisosomes has no effect on the number of eisosomes in cells, related to Figure 4.4. The average number of Fhn1-mCherry-labeled eisosomes per μm2 was measured from the maximum intensity projections of 3 Z-sections through the top surface of 3 randomly selected cells for each of the strains combining Fhn1-mCherry and mGFP-tagged wild type or mutant Myo1. The error bars represent S.D.
201 Chapter 4 Myo1e mutants in yeast
Figure S4.4. Chaperon Rng3 is recruited to the G308R but not the G483D
Myo1 mutant detected using Cam2-mCherry, related to Figure 4.6.
202 Chapter 4 Myo1e mutants in yeast
Figure S4.4. Chaperon Rng3 is recruited to the G308R but not the G483D
Myo1 mutant detected using Cam2-mCherry, related to Figure 4.6. (A-C’’)
Confocal images show the localization of (left panels, A-C) mGFP-tagged mutant
Myo1, (middle panels, A’-C’) mCherry-tagged Cam2 (Cam2-mCherry), and (right panels, A’’-C’’) merge of Myo1-mGFP and Cam2-mCherry images. Cam2- mCherry co-localizes with mGFP-tagged G483D mutant Myo1 in patches (yellow arrowheads), co-localizes with mGFP-tagged G308R Myo1 mutants in eisosomes (yellow arrows), and does not co-localize with GFP-tagged Myo1 mutant lacking the second IQ motif (ΔIQ2) in patches (white arrowheads in C-
C’’). In the ΔIQ2 and, occasionally, in the G308R mutant, Cam2 localizes to cytoplasmic puncta that are devoid of Myo1 (white arrows). (D-F’’) Confocal images show the localization of (left panels, D-F) triple GFP-tagged Rng3 (Rng3-
3xGFP), (middle panels, D’-F’) mCherry-tagged Cam2 (Cam2-mCherry), and
(right panels D’’-F’’) merge of Rng3-3xGFP and Cam2-mCherry images in yeast strains expressing untagged wild type or mutant Myo1 variants. In the G483D mutant, Rng3 localizes to the contractile rings (green arrows) and cytoplasmic spots of unknown nature (green arrowheads in D-D’’) that are distinct from Cam2 in actin patches (red arrowheads). Rng3 co-localizes with Cam2 in eisosomes
(yellow arrows in E-E’’) in the G308R mutant and does not co-localize with Cam2 in eisosomes (red arrows in F’-F’’) in the mutant with duplicated IQ motifs
(2xIQ1IQ2). Cam2 and Rng3 also co-localize in cortical puncta (yellow arrowheads, E-E’’) in the G308R mutant. The images represent maximum
203 Chapter 4 Myo1e mutants in yeast intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4 µm intervals. Scale bar, 10 µm.
204 Chapter 4 Myo1e mutants in yeast
Figure S4.5. Cam2 co-localizes with Myo1 in patches and eisosomes, related to Figure 4.6A-C’’ and Figure S4.4A-C’’.
205 Chapter 4 Myo1e mutants in yeast
Figure S4.5. Cam2 co-localizes with Myo1 in patches and eisosomes, related to Figure 4.6A-C’’ and Figure S4.4A-C’’. Fractions of Myo1 patches and eisosomes that also contained Cam2 and fraction of Cam2 patches/puncta that also contained Myo1 were measured from the maximum intensity projections of 3 Z-sections through the top surface of 3 randomly selected cells for each of the strains combining Cam2-mCherry and mGFP-tagged wild type or mutant
Myo1. The numbers of the structures quantified are indicated in parentheses
206 Chapter 4 Myo1e mutants in yeast
Figure S4.6. Chaperone Rng3 is recruited to the G308R but not the G483D
Myo1 mutant, related to Figure 4.7.
207 Chapter 4 Myo1e mutants in yeast
Figure S4.6. Chaperone Rng3 is recruited to the G308R but not the G483D
Myo1 mutant, related to Figure 4.7. (A-B’’) Confocal images show the localization of (left panels, A-B) triple GFP-tagged Rng3 (Rng3-3xGFP), (middle panels, A’-B’) mCherry-tagged Myo1 variants (Myo1-mCherry), and (right panels
A’’-B’’) merge of Rng3-3xGFP and Myo1-mCherry images. In the G483D mutant,
Rng3 localizes to the contractile rings (green arrows) and cytoplasmic spots of unknown nature (green arrowheads) that are distinct from Myo1(G483D) in actin patches (red arrowheads). In the G308R mutant, Rng3 co-localizes with
Myo1(G308R) in puncta (yellow arrowheads) and eisosomes (yellow arrows). (C-
E’’) Confocal images show the localization of (left panels, C-E) mGFP-tagged
Myo1 variants (Myo1-mGFP), (middle panels, C’-E’) mCherry-tagged Rng3
(Rng3-mCherry), and (right panels C’’-E’’) merge of Myo1-mGFP and Rng3- mCherry images. Rng3 is recruited to Myo1(G308R) in eisosomes (yellow arrows) but not to Myo1(G483D) in patches (green arrowhead) or Myo1 with duplicated IQ motifs (2xIQ1IQ2) in eisosomes (green arrow). The images represent maximum intensity projections of three consecutive optical sections through the top surface of the cell acquired at 0.4 µm interval. Scale bar, 10 µm.
208 Chapter 5 Discussion and Future Directions
Chapter 5
Discussion and Future Directions
209 Chapter 5 Discussion and Future Directions
5.1 Study of Myo1e domain localization and activity at junctions
Myosins are actin-based molecular motors that exhibit strong actin binding in the ADP-bound state and weak actin binding in the ATP-bound state. Myo1e belongs to the Class I myosin. The motor domain of Myo1e is responsible for actin binding and ATPase activity, but has a weaker ATPase activity compared to other conventional myosins (El Mezgueldi et al., 2002). Its tail domain contains three tail homology (TH) subdomains (Fig. 5.1). The TH1 domain is rich in basic residues and binds to negatively charged plasma membrane with high affinity
(Feeser et al., 2010). The TH2 domain is rich in proline residues and may potentially interact with Src homology 3 (SH3) domain-containing proteins. In addition, previous studies have shown that the TH2 domain contains a second actin binding site in some Class I myosins (Jung and Hammer III, 1994). Our data also confirmed that the TH2 domain of Myo1e interacts with actin (See AP
Fig. 2.7). The very C-terminal region of Myo1e is a SH3 domain, which is found in various proteins that are involved in signal transduction and cytoskeleton organization (Musacchio et al., 1992). In mammalian cells, the Myo1e SH3 domain interacts with dynamin and synaptojanin-1, both of which play a role in regulating clathrin-mediated endocytosis (Krendel et al., 2007; Simpson et al.,
1999), and with N-WASP, a regulator of actin nucleation (Cheng et al., 2012, and unpublished data). The Myo1e SH3 domain has been shown to interact with
CARMIL (capping protein, Arp2/3 complex, and Myosin-I linker family proteins) at the leading edge of the cell, playing a role in cell migration (Xu et al., 1995; Liang
210 Chapter 5 Discussion and Future Directions et al., 2009). Our lab has also shown that the SH3 domain of Myo1e interacts with ZO-1, a tight junction associated protein that is also present in the slit diaphragm, via its C-terminus (Bi et al., 2013). In budding yeast Saccharomyces cerevisiae (S. cerevisiae), SH3 domain-containing myosins (Myo3/5p) are associated with proteins that regulate actin assembly (Musacchio et al., 1992), such as Bee1p and verprolin (Vrp1p), which are budding yeast homologs of mammalian WASP (Wiskott-Aldrich syndrome protein) and WIP (WASP- interacting protein), respectively (Anderson et al., 1998; Evangelista et al., 2000;
Geli et al., 2000). In fission yeast S. pombe, SH3 domain containing Myo1 interacts with proline-rich Vrp1p, playing a role in actin assembly at the patches
(Sirotkin et al., 2005). We have also discovered that the SH3 domain of Myo1e interacts with its own TH2 domain in in vitro pulldown assays (data not shown).
Using MDCK cells, we were able to map Myo1e domain localization at cell-cell junctions (Fig. 5.2) (Bi et al., 2013). The tail domain alone (Myo1e-
TH1TH2SH3) is sufficient for targeting Myo1e to cell-cell junctions, plasma membranes, and endocytic vesicles in MDCK cells, suggesting that motor activity is not required in the localization process. The TH2 domain deletion mutants
(Myo1e-ΔTH2, Myo1e-ΔTH2SH3) and the TH2 domain alone (Myo1e-TH2) displayed a diffuse localization in the cytosol rather than a targeted localization to cell-cell junctions, indicating that the TH2 domain of Myo1e is necessary but not sufficient for Myo1e localization. Addition of the TH1 domain (Myo1e-TH1TH2) rescued the diffused phenotype and re-targeted Myo1e protein to cell-cell
211 Chapter 5 Discussion and Future Directions junctions. None of the single domains (Myo1e-TH1, TH2, or SH3 domain) were enriched in cell-cell junctions, likely due to the lack of the other functional domains. This suggests that TH1TH2 is the minimal length of the Myo1e tail domain that is required for its localization.
The motor domain mutants, T119I and A159P, disrupt Myo1e actin- binding ability and ATP hydrolysis. We think that direct disruption of the motor domain of Myo1e destabilizes the protein and making it non-functional. This would explain why these loss-of-function proteins seem diffuse among the cytosol. Our rigor mutant (N164A) remains bound to actin in an ATP-insensitive manner, and has a prominent accumulation at the plasma membrane as we grossly examined the confocal images (See AP Fig. 2.1). We introduced the
N164A mutation in the motor domain of Myo1e-ΔTH2, attempting to rescue the protein distribution from being diffuse in the cytoplasm to being targeted to cell- cell membranes/junctions. However, the Myo1e-N164A- ΔTH2 failed to rescue the phenotype of Myo1e-ΔTH2 (See AP Fig. 2.6), supporting the fact that the
TH2 domain is essential for targeting Myo1e to its cellular location, such as cell- cell contacts.
Class I myosins have been proposed to regulate tension at the plasma membrane, and motor activity is required in this process (Mcconnell and Tyska,
2011; Nambiar et al., 2009). Myo1e is a low duty ratio motor and only stays bound to actin for a very short amount of time (El Mezgueldi et al., 2002).
However, in cell regions rich in actin filaments, Myo1e might become a
212 Chapter 5 Discussion and Future Directions moderate/high duty ratio motor and stay bound to actin for a longer time (Laakso et al., 2008; White et al., 1997). Under this scenario, when the local actin concentration decreases, Myo1e would detach from the actin filaments. Actin polymerization underneath the plasma membrane maintains tension across the plasma membrane and we propose that Myo1e plays a role in anchoring the actin filaments, when they are at a high enough concentration, to the junctions, stabilizing them and thus maintaining membrane tension. Stable junctions have a low or slow exchange rate of junctional components. Non-functional Myo1e would decrease the stability of the junctions and increase turnover. Consistent with this model, our ZO-1 FRAP data showed a higher mobile fraction of ZO-1 in cells expressing the mutant A159P compared to cells expressing wild type
Myo1e. Decreasing junctional stability would decrease membrane tension as actin filament disassembles. Therefore, Myo1e could play a role in regulating tension at the plasma membrane by anchoring the actin filaments to the junctions, and subsequently stabilizing the junctions.
5.2 Model of Myo1e structural organization
Biochemical analysis of the F-actin activated ATPase activity of Myr3, a rat homolog of human Myo1e, showed that inhibition of the tail domain or the truncation of the C-terminal tail of 10kD fragment increased Mg2+-ATPase activity of Myr3, suggesting that the tail domain of Myr3 has an inhibitory effect on the motor domain ATPase activity (Stöffler and Bähler, 1998). One of the possible explanations is that there exists an intramolecular interaction between the very C-
213 Chapter 5 Discussion and Future Directions terminus tail domain and motor domain such that the tail domain folds back on to the motor domain, inhibiting its activity. Another group of scientists discovered a similar but distinct phenomenon when they investigated a Class I myosin in
Acanthamoeba, Myosin IC (AMIC), which is also a homolog of human Myo1e. By using cryo-electron microscopy, X-ray crystallography, and NMR spectroscopy to directly study the crystal structure of the tail domain of AMIC, they discovered that the SH3 domain of AMIC intramolecularly interacted with the pleckstrin homology (PH) domain within the basic region in the AMIC tail (Hwang et al.,
2007; Ishikawa et al., 2004). Truncation analysis of Myo1e domain activity in
MDCK cells indicated that the tail domain alone was twice as enriched at the junctions compared to the full length Myo1e (See Fig. 2.6). This suggested that the Myo1e motor domain (or IQ motif) negatively controlled Myo1e activity (at least localization) at the junctions. A similar result was discovered in Myo1a (a human myosin I isoform). Using single molecule TIRF (total internal reflection fluorescence microscopy), they found that the TH1 domain alone of Myo1a
(Myo1a-TH1) exhibited higher motility than the full-length Myo1a (Myo1a-FL), indicating the inhibitory role of the motor domain to the tail (Mazerik et al., 2014).
In addition to class I myosins, this intramolecular interaction has also been observed in class VII myosin. Umeki et al. observed a decreased ATPase activity in Drosophila Myosin VIIA (DM7A), a homolog of mammalian Myosin VIIA, caused by the tail domain folding back to the head-neck region (Umeki et al.,
2009).
214 Chapter 5 Discussion and Future Directions
Based on the homolog examples we have hypothesized that there might be an intramolecular interaction within the tail domain of Myo1e, possibly between the TH2 domain and SH3 domain, which likely results in a less active form of myosin (Fig. 5.3). However, the TH1 domain is likely exposed, so it can still facilitate Myo1e localizing to the junctions. We think that the intramolecular interaction between the TH2 and SH3 domain is likely weaker than regular protein-protein interactions. Thus, upon the SH3 domain binding to other partners
(such as dynamin, synaptojanin-1, N-WASP, and ZO-1) with higher affinity,
Myo1e opens up and becomes more active. We speculate that Myo1e exists as
TH2-SH3 autoregulatory conformation in the cells until it has been targeted to a cell region with a high concentration of proline-rich containing proteins. Then the
Myo1e SH3 domain binds to them in a higher affinity, which results in the disruption of the intramolecular interaction between the TH2 domain. Myo1e opens up to its more active structure, allowing the TH2 domain binding to actin.
This hypothesis postulates that the switch-off point is coupled to the decreased concentration of SH3 domain binding partners in the cell region, allowing the TH2 domain and the SH3 domain to bind each other without competition and thus deactivating Myo1e. This model of Myo1e activation is presumably located in a cellular region that is rich in actin filaments and proline-rich containing proteins, such as the site of the endocytic vesicle assembly and cell-cell contacts. To test this model, we should perform some biochemical assays, such as analytical ultracentrifugation or electron microscopy to assess the structural conformation of
215 Chapter 5 Discussion and Future Directions
Myo1e. We should also conduct competition binding assays with F-actin using the tail domain alone (TH1+TH2+SH3) or the tail domain without the SH3 domain
(TH1+TH2) to further confirm whether there is a competition interaction to the
Myo1e TH2 domain between F-actin and the Myo1e SH3 domain.
5.3 Potential role of Myo1e in endocytosis – what have we learned from mouse model studies?
Ultrastructure studies in the Myo1e global knockout (KO) mouse model and podocyte-specific KO model have revealed that other than podocyte foot processes effacement and loss of endothelial fenestrations, loss of Myo1e also induced the thickening of the glomerular basement membrane (GBM), which is a classic phenotype usually found in Alport syndrome patients (Chase et al., 2012;
Krendel et al., 2009). Alport syndrome is caused by the mutation in collagen IV genes COL4A3, COL4A4, and COL4A5, which are all major components of mature GBM (Gubler, 2008). After a literature search, we compared the ultrastructure of the GBM in mouse models that are deficient of certain glomerular genes (Table 5.1). Surprisingly, none of the podocyte proteins that are involved in actin filament assembly are associated with defects in GBM integrity. We discovered that the proteins that are associated with GBM thickening in knockouts fall into two groups (highlighted in yellow in Table 5.1). One, such as endophilin, dynamin and synaptojanin, are involved in endocytosis, and the others, such as integrins and integrin binding proteins, collagen IV, laminin play a role in podocytes and GMB adhesion (cell to substrate adhesion). The only slit
216 Chapter 5 Discussion and Future Directions diaphragm associated proteins, when knocked out or knocked in, cause GBM thickening are ZO-1 and nephrin, respectively (Itoh et al., 2014, and unpublished data from Dr. Jones' lab). We have demonstrated that the Myo1e SH3 domain interacts with ZO-1 through in vitro pulldown assays and live-cell imaging (Bi et al., 2013). Our unpublished observations indicated that the Myo1e tail domain binds to chimeric CD8-nephrin in in vitro pulldown assays and associates with nephrin intracellular domain containing vesicles in COS-7 cells. Moreover, Itoh et al. showed that ZO-1, nephrin, and podocin are present in the same complex in co-immunoprecipitation (co-IP) assays. This suggests a similar cellular role or pathway among Myo1e, ZO-1, and nephrin in endocytosis and cell-matrix adhesion.
The same phenotype of GBM thickening in ultrastructure has been found in both integrin-deficient mice kidneys and ZO-1 KO and mutant nephrin knockin mouse models. We speculate that Myo1e may play a role in linking the signaling from the slit diaphragm to the regulation of the basolateral compartment of podocytes, which in turn affects the podocyte adhesion and GBM integrity. Our literature search also revealed that a podocyte protein known as integrin-linked kinase (ILK), which binds to β1 and β3 subunits of integrin, and other adaptor proteins, linking the extracellular matrix to the actin cytoskeleton (Wu and Dedhar,
2001). A recent study using podocyte-specific ILK KO mice revealed that mice without ILK developed proteinuria and the thickening of the GBM upon electron microscopic examinations (Dai et al., 2006). Furthermore, using biochemical
217 Chapter 5 Discussion and Future Directions assays, they showed that ILK associates with nephrin and α – actinin 4 in the same complex, and loss of ILK re-distributed nephrin from the slit diaphragm to the cell body. Therefore, ILK could also be involved in the same process as
Myo1e.
Our literature search suggests that disruption of endocytosis also leads to defects in GBM, likely through integrin signaling. Integrins are transmembrane glycoproteins located at the basal lateral side of podocytes. They exist as heterodimers, containing non-covalently associated α and β subunits. Each unit has a rather large ectodomain, a single transmembrane domain, and a short cytoplasmic tail (Campbell and Humphries, 2011). The activation of integrins is so called “bidirectional”. The “inside-out” activation refers to when an intracellular ligand binds to the cytoplasmic tail of the integrin subunit and induces a conformational change, which in turn activates the ectodomain for extracellular ligand (extracellular matrix) binding. This type of activation is specific for the tight binding to ECM, and is essential for cell-ECM adhesion and cell migration. The
“outside-in” model refers to the extracellular multivalent ligand binding to integrin ectodomain and induces integrin clustering, and subsequently promotes binding to intracellular ligand to the cytoplasmic tail (Shattil et al., 2010). Endocytosis of integrins can be regulated both by the extracellular ligand binding and intracellular cues, and mediated by integrins themselves. If the podocyte endocytic pathway is disrupted, integrin signaling could easily malfunction,
218 Chapter 5 Discussion and Future Directions subsequently leading to GBM defects and podocyte actin cytoskeleton disorganization.
The dominant integrin subunits expressed in kidney podocytes are integrin
α3β1(Pozzi and Zent, 2013), whose ectodomains have high affinity to laminin in
GBM. This strong binding of integrin α3β1 to laminin is mediated by CD151, a tetraspanin family protein that specifically binds to α3 subunit of integrin, but not due to an increase in expression or activation of the integrin (Sachs et al., 2012).
This indicates that loss of podocyte adherens to GBM is due to decreased binding affinity of integrin α3β1 to laminin, which is specifically mediated by
CD151 binding to integrin. Collagen IV specific-binding integrin α1β1 and α2β1 are also expressed in the kidney podocytes. Especially, integrin α1β1 binds to ligand collagen IV and regulates collagen synthesis at the transcription level
(Gardner et al., 1999). The thickening of the GBM observed under the electron microscope might result from the mis-regulation of GBM protein synthesis at the diseased stage.
We propose that in podocytes, Myo1e is involved in the process of endocytosis of nephrin and integrins at podocytes slit diaphragm and basolateral junctions. Disruption of Myo1e might affect the efficiency or accuracy of the endocytic pathway, which in turn might slow down or inhibit endocytosis or exocytosis of transmembrane signaling molecules, such as nephrin and integrin.
Defects in membrane distributions of nephrin and integrin subsequently lead to the disruption of the slit diaphragm and podocyte adhesion, respectively,
219 Chapter 5 Discussion and Future Directions eventually resulting in proteinuria and podocyte deletion or foot process effacement.
Proteinuria is defined as a classic clinical feature of FSGS or nephrotic syndrome. The ultrastructure examination of diseased kidney sections often shows the effacement of podocyte foot processes, and/or defects in GBM and endothelial cells. Obviously, disruption or mutation of matrix proteins is likely to cause the abnormalities in GBM structure, such as the thickening of the GBM.
However, for the genes that do not encode for matrix proteins, the abnormalities in GBM may be secondary to pathophysiology of the disrupted signaling (Miner,
1999). We speculate that the thickening of the GBM is a defensive mechanism of already injured glomerular filtration barrier such that it will slow down further catastrophic damages in the kidney.
5.4 Conclusions
The main goal of this research was to investigate the role of Myosin 1e
(Myo1e) in regulating and maintaining the integrity of cell-cell contacts in mouse kidney podocytes. Our lab has shown that disruption of Myo1e in mice promoted kidney injury, characterized by proteinuria, podocyte foot process effacement and the thickening of the glomerular basement membrane (Chase et al., 2012;
Krendel et al., 2009). Furthermore, genetic studies by different groups identified several Myo1e mutations that are associated with focal segmental glomerulosclerosis, the defining feature of which is proteinuria (Al-Hamed et al.,
220 Chapter 5 Discussion and Future Directions
2013; Mele et al., 2011; Sanna-Cherchi et al., 2011). Using biochemical assays and image analysis, we demonstrated that Myo1e is a component of the renal slit diaphragm, and without functional Myo1e, the nascent junction assembly is defective (Bi et al., 2013). In the future, detailed and accurate analysis should be performed by examining the assembly of the cell-cell junctions, and the recruitment of junctional transmembrane components, such as nephrin or cadherins. We proposed that the recycling of nephrin is likely Myo1e-mediated.
Besides cell culture systems, we also used fission yeast as a versatile tool to characterize two kidney disease associated mutants and discover that both of the mutants are non-functional, but in different ways (Bi et al., 2015). Further studies might also use human Myo1e to replace fission yeast Myo1, as it would be interesting to see whether human Myo1e rescues any defects that result from deletion of myo1 gene in yeast.
Based on the clinical studies and our own investigations, we demonstrate that Myo1e is important for kidney function in higher vertebrates. In the future, it would be beneficial to explore the underlying mechanism linking Myo1e to
FSGS/nephrotic syndrome, which will shine some light on the pathogenesis of primary kidney disease at the molecular level.
221 Chapter 5 Discussion and Future Directions
5.5 Reference:
Abrahamson, D.R., Isom, K., Roach, E., Stroganova, L., Zelenchuk, A., Miner, J.H., St John, P.L., 2007. Laminin compensation in collagen alpha3(IV) knockout (Alport) glomeruli contributes to permeability defects. J. Am. Soc. Nephrol. 18, 2465–2472. doi:10.1681/ASN.2007030328 Abrass, C.K., Hansen, K.M., Patton, B.L., 2010. Laminin alpha4-null mutant mice develop chronic kidney disease with persistent overexpression of platelet-derived growth factor. Am. J. Pathol. 176, 839–849. doi:10.2353/ajpath.2010.090570 Akiyama, K., Morita, H., Suetsugu, S., Kuraba, S., Numata, Y., Yamamoto, Y., Inui, K., Ideura, T., Wakisaka, N., Nakano, K., Oniki, H., Takenawa, T., Matsuyama, M., Yoshimura, A., 2008. Actin -related protein 3 (Arp3) is mutated in proteinuric BUF/Mna rats. Mamm. Genome 19, 41–50. doi:10.1007/s00335-007-9078-5 Al-Hamed, M.H., Al-Sabban, E., Al-Mojalli, H., Al-Harbi, N., Faqeih, E., Al Shaya, H., Alhasan, K., Al-Hissi, S., Rajab, M., Edwards, N., Al-Abbad, A., Al-Hassoun, I., Sayer, J. a, Meyer, B.F., 2013. A molecular genetic analysis of childhood nephrotic syndrome in a cohort of Saudi Arabian families. J. Hum. Genet. 1–10. doi:10.1038/jhg.2013.27 Anderson, B.L., Boldogh, I., Evangelista, M., Boone, C., Greene, L. A., Pon, L. A, 1998. The Src homology domain 3 (SH3) of a yeast type I myosin, Myo5p, binds to verprolin and is required for targeting to sites of actin polarization. J. Cell Biol. 141, 1357–1370. doi:10.1083/jcb.141.6.1357 Asanuma, K., Kim, K., Oh, J., Giardino, L., Chabanis, S., Faul, C., Reiser, J., Mundel, P., 2005. Synaptopodin regulates the actin-bundling activity of α-actinin in an isoform- specific manner. J. Clin. Invest. 115, 1188–98. doi:10.1172/JCI200523371.1188 Asanuma, K., Yanagida-Asanuma, E., Faul, C., Tomino, Y., Kim, K., Mundel, P., 2006. Synaptopodin orchestrates actin organization and cell motility via regulation of RhoA signalling. Nat. Cell Biol. 8, 485–91. doi:10.1038/ncb1400 Bi, J., Carroll, R.T., James, M.L., Ouderkirk, J.L., Krendel, M., Sirotkin, V., 2015. Effects of FSGS-associated mutations on the stability and function of myosin-1 in fission yeast. Dis. Model. Mech. 8(8):891-902. doi:10.1242/dmm.020214 Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M., 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am. J. Physiol. Renal Physiol. 305, 532–544. doi:10.1152/ajprenal.00223.2013 Bisson, N., Ruston, J., Jeansson, M., Vanderlaan, R., Hardy, W.R., Du, J., Hussein, S.M., Coward, R.J., Quaggin, S.E., Pawson, T., 2012. The Adaptor Protein Grb2 Is Not Essential for the Establishment of the Glomerular Filtration Barrier. PLoS One 7. doi:10.1371/journal.pone.0050996 Buday, L., Wunderlich, L., Tamás, P., 2002. The Nck family of adapter proteins: Regulators of actin cytoskeleton. Cell. Signal. 14, 723–731. doi:10.1016/S0898-6568(02)00027-X Campbell, I.D., Humphries, M.J., 2011. Integrin structure, activation, and interactions. Cold Spring Harb. Perspect. Biol. 3, 1–14. doi:10.1101/cshperspect.a004994 Chase, S.E., Encina, C. V, Stolzenburg, L.R., Tatum, A.H., Holzman, L.B., Krendel, M., 2012. Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am. J. Physiol. Renal Physiol. 303, F1099–106. doi:10.1152/ajprenal.00251.2012 Cheng, J., Grassart, A., Drubin, D.G., 2012. Myosin 1E coordinates actin assembly and cargo trafficking during clathrin-mediated endocytosis. J. Cell Sci. 23(15):2891-904. Cosgrove, D., Meehan, D.T., Grunkemeyer, J. a., Kornak, J.M., Sayers, R., Hunter, W.J., Samuelson, G.C., 1996. Collagen COL4A3 knockout: A mouse model for autosomal Alport syndrome. Genes Dev. 10, 2981–2992. doi:10.1101/gad.10.23.2981
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Dai, C., Stolz, D.B., Bastacky, S.I., St-Arnaud, R., Wu, C., Dedhar, S., Liu, Y., 2006. Essential role of integrin-linked kinase in podocyte biology: Bridging the integrin and slit diaphragm signaling. J. Am. Soc. Nephrol. 17, 2164–2175. doi:10.1681/ASN.2006010033 Donoviel, D.B., Freed, D.D., Vogel, H., David, G., Hawkins, E., Barrish, J.P., Mathur, B.N., Turner, A., Geske, R., Montgomery, C. a, Starbuck, M., Brandt, M., Gupta, A., Zambrowicz, B.P., Powell, D.R., Potter, D.G., Turner, C.A., Ramirez-solis, R., 2001. Proteinuria and Perinatal Lethality in Mice Lacking NEPH1 , a Novel Protein with Homology to NEPHRIN Proteinuria and Perinatal Lethality in Mice Lacking NEPH1 , a Novel Protein with Homology to NEPHRIN. Mol. Cell. Biol. 21, 4829–36. doi:10.1128/MCB.21.14.4829 El Mezgueldi, M., Tang, N., Rosenfeld, S.S., Ostap, E.M., 2002. The kinetic mechanism of Myo1e (human myosin-IC). J. Biol. Chem. 277, 21514–21. doi:10.1074/jbc.M200713200 Evangelista, M., Klebl, B.M., Tong, A.H.Y., Webb, B. a., Leeuw, T., Leberer, E., Whiteway, M., Thomas, D.Y., Boone, C., 2000. A role for myosin-I in actin assembly through interactions with Vrp1p, Bee1p, and the Arp2/3 complex. J. Cell Biol. 148, 353–362. doi:10.1083/jcb.148.2.353 Feeser, E. a, Ignacio, C.M.G., Krendel, M., Ostap, E.M., 2010. Myo1e binds anionic phospholipids with high affinity. Biochemistry 49, 9353–60. doi:10.1021/bi1012657 Gardner, H., Broberg, a, Pozzi, a, Laato, M., Heino, J., 1999. Absence of integrin alpha1beta1 in the mouse causes loss of feedback regulation of collagen synthesis in normal and wounded dermis. J. Cell Sci. 112 ( Pt 3, 263–272. Geli, M.I., Lombardi, R., Schmelzl, B., Riezman, H., 2000. An intact SH3 domain is required for myosin I-induced actin polymerization. EMBO J. 19, 4281–4291. doi:10.1093/emboj/19.16.4281 Georges-Labouesse, E., Messaddeq, N., Yehia, G., Cadalbert, L., Dierich, a, Le Meur, M., 1996. Absence of integrin alpha 6 leads to epidermolysis bullosa and neonatal death in mice. Nat. Genet. 13, 370–373. doi:10.1038/ng0796-370 Goley, E.D., Welch, M.D., 2006. The ARP2/3 complex: an actin nucleator comes of age. Nat. Rev. Mol. Cell Biol. 7, 713–726. doi:10.1038/nrm2026 Gubler, M. C., 2008. Inherited disease of the glomerular basement membrane. Nat. Clin. Pract. Nephrol. 4, 24–37. Harita, Y., Kurihara, H., Kosako, H., Tezuka, T., Sekine, T., Igarashi, T., Hattori, S., 2008. Neph1, a component of the kidney slit diaphragm, is tyrosine-phosphorylated by the Src family tyrosine kinase and modulates intracellular signaling by binding to Grb2. J. Biol. Chem. 283, 9177–9186. doi:10.1074/jbc.M707247200 Heino, J., 2000. The collagen receptor integrins have distinct ligand recognition and signaling functions. Matrix Biol. 19, 319–323. doi:10.1016/S0945-053X(00)00076-7 Hinamoto, N., Maeshima, Y., Yamasaki, H., Nasu, T., Saito, D., Watatani, H., Ujike, H., Tanabe, K., Masuda, K., Arata, Y., Sugiyama, H., Sato, Y., Makino, H., 2014. Exacerbation of Diabetic Renal Alterations in Mice Lacking Vasohibin-1. PLoS One 9, e107934. doi:10.1371/journal.pone.0107934 Huber, T.B., Kwoh, C., Wu, H., Asanuma, K., Godel, M., Hartleben, B., Blumer, K.J., Miner, J.H., Mundel, P., Shaw, A.S., 2006. Bigenic mouse models of focal segmental glomerulosclerosis involving pairwise interaction of CD2AP, Fyn, and synaptopodin. J. Clin. Invest. 116, 1337–1345. doi:10.1172/JCI27400.to Hwang, K.-J., Mahmoodian, F., Ferretti, J. a, Korn, E.D., Gruschus, J.M., 2007. Intramolecular interaction in the tail of Acanthamoeba myosin IC between the SH3 domain and a putative pleckstrin homology domain. Proc. Natl. Acad. Sci. U. S. A. 104, 784–9. doi:10.1073/pnas.0610231104 Ishikawa, T., Cheng, N., Liu, X., Korn, E.D., Steven, A.C., 2004. Subdomain organization of the Acanthamoeba myosin IC tail from cryo-electron microscopy. Proc. Natl. Acad. Sci. U. S. A. 101, 12189–94. doi:10.1073/pnas.0404835101
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Itoh, M., Nakadate, K., Horibata, Y., Matsusaka, T., Xu, J., Hunziker, W., Sugimoto, H., 2014. The Structural and Functional Organization of the Podocyte Filtration Slits Is Regulated by Tjp1/ZO-1. PLoS One 9, e106621. doi:10.1371/journal.pone.0106621 Jones, N., Blasutig, I.M., Eremina, V., Ruston, J.M., Bladt, F., Li, H., Huang, H., Larose, L., Li, S.S.-C., Takano, T., Quaggin, S.E., Pawson, T., 2006. Nck adaptor proteins link nephrin to the actin cytoskeleton of kidney podocytes. Nature 440, 818–823. doi:10.1038/nature04662 Jones, N., New, L. a, Fortino, M. a, Eremina, V., Ruston, J., Blasutig, I.M., Aoudjit, L., Zou, Y., Liu, X., Yu, G.-L., Takano, T., Quaggin, S.E., Pawson, T., 2009. Nck proteins maintain the adult glomerular filtration barrier. J. Am. Soc. Nephrol. 20, 1533–43. doi:10.1681/ASN.2009010056 Jung, G., Hammer III, J.A., 1994. The actin binding site in the tail domain of Dictyostelium myosin IC ( tail homology region 2 ). FEBS Lett. 342, 197–202. Kanasaki, K., Kanda, Y., Palmsten, K., Tanjore, H., Lee, S.B., Lebleu, V.S., Gattone Jr., V.H., Kalluri, R., 2008. Integrin beta1 mediated matrix assembly and signaling is critical for the normal development and function of the kidney glomerulus. Dev. Biol. 313, 584–593. doi:10.1016/j.biotechadv.2011.08.021.Secreted Krendel, M., Kim, S. V, Willinger, T., Wang, T., Kashgarian, M., Flavell, R. a, Mooseker, M.S., 2009. Disruption of Myosin 1e promotes podocyte injury. J. Am. Soc. Nephrol. 20, 86–94. doi:10.1681/ASN.2007111172 Krendel, M., Osterweil, E.K., Mooseker, M.S., 2007. Myosin 1E interacts with synaptojanin-1 and dynamin and is involved in endocytosis. FEBS Lett. 581, 644–50. doi:10.1016/j.febslet.2007.01.021 Laakso, J.M., Lewis, J.H., Shuman, H., Ostap, E.M., 2008. Myosin-I can act as a molecular force sensor. Science (80). 321, 133–136. doi:10.1016/j.biotechadv.2011.08.021.Secreted Le Clainche, C., Carlier, M. F., 2008. Regulation of actin assembly associated with protrusion and adhesion in cell migration. Physiol. Rev. 88, 489–513. doi:10.1152/physrev.00021.2007 Liang, Y., Niedeerstrasser, H., Edwards, M., Jackson, C.E., Cooper, J.A., 2009. Distinct roles for CARMIL isofrms in cell migration. Mol. Biol. Cell 20, 5290–5305. doi:10.1091/mbc.E08 Liu, G., Kaw, B., Kurfis, J., Rahmanuddin, S., Kanwar, Y.S., Chugh, S.S., 2003. Neph1 and nephrin interaction in the slit diaphragm is an important determinant of glomerular permeability. J. Clin. Invest. 112, 209–221. doi:10.1172/JCI200318242 Liu, X.L., Kilpeläinen, P., Hellman, U., Sun, Y., Wartiovaara, J., Morgunova, E., Pikkarainen, T., Yan, K., Jonsson, A.P., Tryggvason, K., 2005. Characterization of the interactions of the nephrin intracellular domain: Evidence that the scaffolding protein IQGAP1 associates with nephrin. FEBS J. 272, 228–243. doi:10.1111/j.1432-1033.2004.04408.x Lu, W., Katz, S., Gupta, R., Mayer, B.J., 1997. Activation of Pak by membrane localization mediated by an SH3 domain from the adaptor protein Nck. Curr. Biol. 7, 85–94. doi:10.1016/S0960-9822(06)00052-2 Mazerik, J.N., Kraft, L.J., Kenworthy, A.K., Tyska, M.J., 2014. Motor and tail homology 1 (TH1) domains antagonistically control myosin-1 dynamics. Biophys. J. 106, 649–658. doi:10.1016/j.bpj.2013.12.038 Mcconnell, R.E., Tyska, M.J., 2011. Leveraging the membrane-cytoskeleton interface with myosin-1. Mol. Biol. Cell 20, 418–426. doi:10.1016/j.tcb.2010.04.004.Leveraging Mele, C., D, B.S., Iatropoulos, P., Donadelli, R., Calabria, A., Eng, D., Maranta, R., Cassis, P., Ph, D., Buelli, S., Tomasoni, S., Piras, R., D, C.P., Krendel, M., Bettoni, S., Biotech, D., Morigi, M., Delledonne, M., Pecoraro, C., Abbate, I., Capobianchi, M.R., Hildebrandt, F., Otto, E., Schaefer, F., Macciardi, F., Ozaltin, F., Emre, S., Ibsirlioglu, T., Benigni, A., Remuzzi, G., 2011. MYO1E mutations and childhood familial focal segmental glomerulosclerosis. N. Engl. J. Med. 365, 295–306.
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Miner, J.H., 1999. Renal basement membrane components. Kidney Int 56, 2016–2024. doi:10.1046/j.1523-1755.1999.00785.x Miner, J.H., Li, C., 2000. Defective glomerulogenesis in the absence of laminin alpha5 demonstrates a developmental role for the kidney glomerular basement membrane. Dev. Biol. 217, 278–289. doi:10.1006/dbio.1999.9546 Morello, R., Zhou, G., Dreyer, S.D., Harvey, S.J., Ninomiya, Y., Thorner, P.S., Miner, J.H., Cole, W., Winterpacht, a, Zabel, B., Oberg, K.C., Lee, B., 2001. Regulation of glomerular basement membrane collagen expression by LMX1B contributes to renal disease in nail patella syndrome. Nat. Genet. 27, 205–208. doi:10.1038/84853 Musacchio, a, Gibson, T., Lehto, V.P., Saraste, M., 1992. SH3--an abundant protein domain in search of a function. FEBS Lett. 307, 55–61. Nambiar, R., McConnell, R.E., Tyska, M.J., 2009. Control of cell membrane tension by myosin-I. Proc. Natl. Acad. Sci. U. S. A. 106, 11972–11977. doi:10.1073/pnas.0901641106 Noakes, P.G., Miner, J.H., Gautam, M., Cunningham, J.M., Sanes, J.R., P., M.J., 1995. The renal glomerulus of mice lacking s-laminin/laminin beta2: nephrosis despite molecular compensation by laminin beta1. Nat. Genet. 10, 400–406. Palmén, T., Lehtonen, S., Ora, A., Kerjaschki, D., Antignac, C., Lehtonen, E., Holthöfer, H., 2002. Interaction of endogenous nephrin and CD2-associated protein in mouse epithelial M-1 cell line. J. Am. Soc. Nephrol. 13, 1766–1772. doi:10.1097/01.ASN.0000019842.50870.41 Pozzi, A., Jarad, G., Moeckel, G.W., Coffa, S., Zhang, X., Gewin, L., Eremina, V., Hudson, B.G., Borza, D. B., Harris, R.C., Holzman, L.B., Phillips, C.L., Fassler, R., Quaggin, S.E., Miner, J.H., Zent, R. 2008. Beta1 integrin expression by podocytes is required to maintain glomerular structural integrity. Dev. Biol. 316, 288–301. doi:10.1016/j.jsbmb.2011.07.002.Identification Pozzi, A., Zent, R., 2013. Integrins in kidney disease. J. Am. Soc. Nephrol. 24, 1034–9. doi:10.1681/ASN.2013010012 Putaala, H., Soininen, R., Kilpeläinen, P., Wartiovaara, J., Tryggvason, K., 2001. The murine nephrin gene is specifically expressed in kidney, brain and pancreas: inactivation of the gene leads to massive proteinuria and neonatal death. Hum. Mol. Genet. 10, 1–8. Rantanen, M., Palmén, T., Pätäri, A., Ahola, H., Lehtonen, S., Aström, E., Floss, T., Vauti, F., Wurst, W., Ruiz, P., Kerjaschki, D., Holthöfer, H., 2002. Nephrin TRAP mice lack slit diaphragms and show fibrotic glomeruli and cystic tubular lesions. J. Am. Soc. Nephrol. 13, 1586–1594. doi:10.1097/01.ASN.0000016142.29721.22 Rohatgi, R., Nollau, P., Henry Ho, H.Y., Kirschner, M.W., Mayer, B.J., 2001. Nck and Phosphatidylinositol 4,5-Bisphosphate Synergistically Activate Actin Polymerization through the N-WASP-Arp2/3 Pathway. J. Biol. Chem. 276, 26448–26452. doi:10.1074/jbc.M103856200 Sachs, N., Claessen, N., Aten, J., Kreft, M., Teske, G.J.D., Koeman, A., Zuurbier, C.J., Janssen, H., Sonnenberg, A., 2012. Blood pressure influences end-stage renal disease of Cd151 knockout mice. Clin. Investig. (Lond). 122, 348–358. doi:10.1172/JCI58878DS1 Sachs, N., Kreft, M., Van Den Bergh Weerman, M. a., Beynon, A.J., Peters, T. a., Weening, J.J., Sonnenberg, A., 2006. Kidney failure in mice lacking the tetraspanin CD151. J. Cell Biol. 175, 33–39. doi:10.1083/jcb.200603073 Sanna-Cherchi, S., Burgess, K.E., Nees, S.N., Caridi, G., Weng, P.L., Dagnino, M., Bodria, M., Carrea, A., Allegretta, M. a, Kim, H.R., Perry, B.J., Gigante, M., Clark, L.N., Kisselev, S., Cusi, D., Gesualdo, L., Allegri, L., Scolari, F., D’Agati, V., Shapiro, L.S., Pecoraro, C., Palomero, T., Ghiggeri, G.M., Gharavi, A.G., 2011. Exome sequencing identified MYO1E and NEIL1 as candidate genes for human autosomal recessive steroid-resistant nephrotic syndrome. Kidney Int. 80, 389–96. doi:10.1038/ki.2011.148 Schell, C., Baumhakl, L., Salou, S., Conzelmann, A. C., Meyer, C., Helmstadter, M., Wrede, C., Grahammer, F., Eimer, S., Kerjaschki, D., Walz, G., Snapper, S., Huber, T.B., 2013. N-WASP Is Required for Stabilization of Podocyte Foot Processes. J. Am. Soc. Nephrol. 24, 713–721. doi:10.1681/ASN.2012080844
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Shattil, S.J., Kim, C., Ginsberg, M.H., 2010. The final steps of integrin activation: the end game. Nat. Rev. Mol. Cell Biol. 11, 288–300. doi:10.1038/nrm2871 Shih, N.Y., Li, J., Karpitskii, V., Nguyen, a, Dustin, M.L., Kanagawa, O., Miner, J.H., Shaw, a S., 1999. Congenital nephrotic syndrome in mice lacking CD2-associated protein. Science 286, 312–315. doi:10.1126/science.286.5438.312 Simpson, F., Hussain, N.K., Qualmann, B., Kelly, R.B., Kay, B.K., McPherson, P.S., Schmid, S.L., 1999. SH3-domain-containing proteins function at distinct steps in clathrin-coated vesicle formation. Nat. Cell Biol. 1, 119–24. doi:10.1038/10091 Sirotkin, V., Beltzner, C.C., Marchand, J.-B., Pollard, T.D., 2005. Interactions of WASp, myosin-I, and verprolin with Arp2/3 complex during actin patch assembly in fission yeast. J. Cell Biol. 170, 637–48. doi:10.1083/jcb.200502053 Soda, K., Balkin, D.M., Ferguson, S.M., Paradise, S., Milosevic, I., Giovedi, S., Volpicelli-Daley, L., Tian, X., Wu, Y., Ma, H., Son, S.H., Zheng, R., Moeckel, G., Cremona, O., Holzman, L.B., De Camilli, P., Ishibe, S., 2012. Role of dynamin, synaptojanin, and endophilin in podocyte foot processes. J. Clin. Invest. 122, 4401–4411. doi:10.1172/JCI65289 Stein, P.L., Vogel, H., Soriano, P., 1994. Combined deficiencies of src, fyn, and yes tyrosine kinases in mutant mice. Genes Dev. 8, 1999–2007. doi:10.1101/gad.8.17.1999 Stöffler, H.E., Bähler, M., 1998. The ATPase activity of Myr3, a rat myosin I, is allosterically inhibited by its own tail domain and by Ca2+ binding to its light chain calmodulin. J. Biol. Chem. 273, 14605–11. Suzuki, Y., Kobayashi, M., Miyashita, H., Ohta, H., Sonoda, H., Sato, Y., 2010. Isolation of a small vasohibin-binding protein (SVBP) and its role in vasohibin secretion. J. Cell Sci. 123, 3094–3101. doi:10.1242/jcs.067538 Umeki, N., Jung, H.S., Watanabe, S., Sakai, T., Li, X., Ikebe, R., Craig, R., Ikebe, M., 2009. The tail binds to the head-neck domain, inhibiting ATPase activity of myosin VIIA. Proc. Natl. Acad. Sci. U. S. A. 106, 8483–8488. doi:10.1073/pnas.0812930106 Verma, R., Wharram, B., Kovari, I., Kunkel, R., Nihalani, D., Wary, K.K., Wiggins, R.C., Killen, P., Holzman, L.B., 2003. Fyn binds to and phosphorylates the kidney slit diaphragm component Nephrin. J. Biol. Chem. 278, 20716–20723. doi:10.1074/jbc.M301689200 White, H.D., Belknap, B., Webb, M.R., 1997. Kinetics of nucleoside triphosphate cleavage and phosphate release steps by associated rabbit skeletal actomyosin, measured using a novel fluorescent probe for phosphate. Biochemistry 36, 11828–11836. doi:10.1021/bi970540h Wu, C., Dedhar, S., 2001. Integrin-linked kinase (ILK) and its interactors: A new paradigm for the coupling of extracellular matrix to actin cytoskeleton and signaling complexes. J. Cell Biol. 155, 505–510. doi:10.1083/jcb.200108077 Xu, P., Zot, a. S., Zot, H.G., 1995. Identification of Acan125 as a myosin-I-binding protein present with myosin-I on cellular organelles of Acanthamoeba. J. Biol. Chem. 270, 25316–25319. doi:10.1074/jbc.270.43.25316 Yanagida-Asanuma, E., Asanuma, K., Kim, K., Donnelly, M., Young Choi, H., Hyung Chang, J., Suetsugu, S., Tomino, Y., Takenawa, T., Faul, C., Mundel, P., 2007. Synaptopodin protects against proteinuria by disrupting Cdc42:IRSp53:Mena signaling complexes in kidney podocytes. Am. J. Pathol. 171, 415–427. doi:10.2353/ajpath.2007.070075 Yu, C.C.K., Yen, T.S.B., Lowell, C. A., DeFranco, A.L., 2001. Lupus-like kidney disease in mice deficient in the Src family tyrosine kinases Lyn and Fyn. Curr. Biol. 11, 34–38. doi:10.1016/S0960-9822(00)00024-5
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Figure 5.1. Cartoon showing Myo1e subdomain interactions.
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Figure 5.1. Cartoon showing Myo1e subdomain interactions. Motor domain of Myo1e (colored in blue) binds to actin in an ATP-dependent manner. The IQ motif (colored in yellow) binds to light chain calmodulin. The TH1 domain (colored in magenta) is positively charged and binds to negatively charged plasma membrane. The TH2 domain (light green) contains proline-rich motifs (PXXP).
The SH3 domain (colored in orange) interacts with proline-rich containing proteins, such as ZO-1, dynamin, synaptojanin-1, and CARMIL.
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Figure 5.2. Summary of junction localization of Myo1e constructs.
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Figure 5.2. Summary of junction localization of Myo1e constructs. Plus signs (+) mean positive localization at junctions, while minus signs (-) mean lack of junction localization. The bolded construct names indicate they are enriched at junctions.
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Figure 5.3. Schematic model of Myo1e structural conformation.
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Figure 5.3. Schematic model of Myo1e structural conformation. Upon the
SH3 domain binding to proline-rich motif containing proteins, Myo1e opens up from a less active form to a more open and active form, allowing effective interactions with F-actin and other binding partners and to carry out cellular activities.
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Table 5.1: Knockout mouse models leading to proteinuria
Protein Knockout Kidney Known Reference conditions ultrastructure interacting defects partner in podocytes Podocyte cytoplasmic proteins Nck Constitutive Absence of foot Nephrin (Buday et al., Nck1-/-, process at N-WASP 2002; Jones Nck2fl/fl E16.5 (fail to PAK et al., 2006; form foot Synaptojanin Lu et al., process), 1 1997; Normal GBM Rohatgi et al., 2001) Podocyte Foot process (Jones et al., specific effacement, 2009) (inducible) collapsed actin KO of Nck2 in along GBM, Nck1-null Normal GBM mice N-WASP Podocyte Electron-dense SH3 domains (Schell et al., specific KO, actin cluster of Nck ½, 2013) and podocyte near GBM, foot Grb2, Fyn, Also see specific process Myo1e1 review (Le inducible KO effacement, Small Clainche and Normal GBM GTPase Carlier, 2008) G-actin Arp2/3 complex PIP2 WIP Arp3 Missense Election-dense Arp2/3 (Akiyama et mutation in actin, complex al., 2008) BUF/Mna Normal GBM N-WASP Also see kidney Actin review (Goley and Welch, 2006) Grb2 Podocyte No significant Neph1 (Bisson et al., specific KO alteration in N-WASP 2012) podocyte (Harita et al., structure 2008) Synaptopodin Constitutive No significant CD2AP (Yanagida- KO of Synpo- alteration in Myo1e2 Asanuma et
233 Chapter 5 Discussion and Future Directions
long and podocyte IRSp53 al., 2007) Synpo-short, structure in α – actinin (Asanuma et but still normal 2/4 al., 2005) express condition, but Active RhoA (Huber et al., Synpo-T in foot process 2006) podocytes remains (Asanuma et effaced upon al., 2006) recovery Normal GBM Synpo+/- and Foot process (Huber et al., Synpo-/- on a effacement, 2006) Cd2ap+/- Normal GBM background Myosin 1e Constitutive FP effacement Dynamin (Bi et al., KO GBM Synaptojanin- 2013; thickening3 1 Krendel et Proximal tubule N-WASP1 al., 2007) lack of ZO-1 (Krendel et microvilli al., 2009) Podocyte (Chase et al., specific KO, 2012; Inducible KO Dynamin-1/2 Conditional FP is normal at Myo1e (Soda et al., KO both early age, but Endophilin-1 2012) Dynamin 1 becomes CD2AP (Krendel et and 2 in effaced at al., 2007) podocytes week 4; Immature endocytic events GBM thickening Synaptojanin- Constitutive FP effacement Myo1e (Soda et al., 1 double KO Immature Clathrin 2012) (die endocytic Endophilin-1 (Krendel et perinatally) events CD2AP al., 2007) GBM thickening Endophilin- Constitutive Normal Dynamin (Soda et al., 1/2/3 triple KO embryonic Synaptojanin- 2012) (die development 1 perinatally) FP effacement CD2AP GBM thickening
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CD2AP Constitutive FP effacement Dynamin (Shih et al., KO Normal GBM Synaptojanin 1999) and endothelial Endophilin (Huber et al., cells Synaptopodin 2006) Massive extracellular matrix deposition in later age (mesangial cells) Integrin-linked Podocyte- Podocyte Nephrin (Dai et al., kinase (ILK) specific KO apoptosis and α – actinin4 2006) detachment Integrin α3β1 GBM thickening Podocyte slit diaphragm associated proteins Nephrin Constitutive FP effacement Nck (Putaala et KO, Absent of the CD2AP al., 2001) Nephrintrap/trap SD Fyn (Rantanen et Normal GBM PI3K al., 2002) IQGAP1 (Liu et al., ZO-1 2005; Palmén et al., 2002) (Verma et al., 2003) C57BL/6 Normal FP, Unpublished NephrinY3F/Y3F GBM and data from Dr. knockin endothelial Nina Jones (lacking Nck cells at 1 lab interaction) month of age; At later age, mild locally effaced FP and GBM thickening CD-1 Progressive NephrinY3F/Y3F disorganized knockin FP, Slightly thickened GBM (irregular GBM “spikes”)
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Fyn Constitutive Mild phenotype Nephrin (Verma et al., KO 2003) (Yu et al., 2001) Fyn-/- yes-/- Expansion of (Stein et al., double KO mesagial 1994) matrix Neph1 Constitutive FP effacement Nephrin (Liu et al., KO ZO-1 2003) Grb2 (Donoviel et Src family al., 2001) tyrosin kinase (Harita et al., (SFK) 2008) ZO-1/Tjp-1 Podocyte Loss of SD Nephrin (Itoh et al., specific KO Disruption of Myo1e 2014) FP (Bi et al., Thickened 2013) GBM Podocytes basal-lateral membrane associated proteins Integrin α3β1 Constitutive Proximal tubule Laminin (Kanasaki et KO α3 lack of CD151 al., 2008) subunit microvilli GBM thickening Absence of podocyte formation Lack of fusion of dual BM, Disorganization and fragmentation of GBM Podocyte- Thickening of (Sachs et al., specific GBM 2006) conditional α3 FP effacement subunit KO Podocyte- FP effacement (Pozzi et al., specific β1 Normal GBM 2008) subunit KO composition in ultrastructure Abnormal capillary loops and the
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mesangium Podocyte- FP effacement (Kanasaki et specific β1 Lamination of al., 2008) subunit KO GBM (thickening) Integrin α6β1 Constitutive No significant Laminin (Georges- KO α6 alteration in Labouesse et subunit glomerular al., 1996) structure
Integrin α6β4 No mouse Laminin model yet Integrin α1β1 Constitutive No obvious Collagen IV (Heino, 2000) α1 subunit renal KO phenotype Integrin α2β1 Constitutive No obvious Collagen I α2 subunit renal KO phenotype CD151 Constitutive Thickening of Integrin α3β1 (Sachs et al., KO GBM 2006) Loss of (Sachs et al., endothelial 2012) fenestrations FP effacement However, can be normal GBM and FP LMX1B (LIM Constitutive GBM COL4A4 (Morello et homeodomain KO thickening and COL4A3 al., 2001) transcription split factor) Glomerular basement membrane (GBM) associated proteins COL4A3 Constitutive GBM LMX1B (Cosgrove et KO thickening and al., 1996) thinning, (Abrahamson multilamination et al., 2007) and splitting FP effacement Laminin β2 Constitutive Normal GBM (Noakes et chain KO al., 1995) Laminin α5 Constitutive Loss of GBM at (Miner and chain KO capillary loop Li, 2000) stage Mesagial cells associated proteins
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Laminin α4 Constitutive Normal GBM, (Abrass et chain KO FP, and al., 2010) endothelial fenestrae Dilated capillary loops Mesangial interposition Endothelial cells apoptosis
Endothelial cells associated proteins Vasohibin-1 Induced GBM Small (Hinamoto et diabetic thickening vasohibin- al., 2014) model in FP effacement binding (Suzuki et al., heterozygous Decrease in protein 2010) VASH1+/- SD density (SVBP) 1, Our unpublished data
2, In in vitro pulldown assays (See AP Fig. 1.3)
3, The phenotype of GBM thickening is highlighted in yellow.
238 Appendix 1 Synaptopodin project
Appendix Chapter 1
Analysis of the interactions between
Synaptopodin and Myosin 1e
Appendix chapter 1 is written by myself and edited by my colleagues for grammatical errors. This chapter is not yet ready for publication, since there are still a few more experiments to perform in order to finish the story.
239 Appendix 1 Synaptopodin project
AP 1.1 Introduction
Synaptopodin (Synpo) is an actin-associated protein that is expressed in differentiated podocytes in the kidney and postsynaptic densities in the brain
(restricted to telencephalic neurons) (Mundel et al., 1991)Mundel et al., 1997).
Sequencing analysis shows that 20% of Synpo’s amino acids are prolines, which makes it a linear structure rather than globular (Mundel et al., 1997). Because
Synpo is rich in proline, it is more susceptible to proteolytic degradation (Mundel et al., 1997). The expression pattern of Synpo is differentiation dependent.
During kidney development, Synpo is first detected at the capillary loop stage, when the primitive podocytes start differentiating (Mundel and Kriz, 1995). Three isoforms of Synpo (Fig. 1A) have been discovered at this time; N-terminal Synpo- short (1-685aa) is expressed in neurons, and full length Synpo-long (1-903aa) and C-terminal Synpo-T (712-903aa) are expressed in kidneys (Asanuma et al.,
2005).
Synpo localizes as puncta, a periodic dotted pattern, along the stress fibers and in focal adhesions. Using an actin depolymerization reagent such as
Cytochalasin B causes the redistribution of Synpo from the stress fibers to cytoplasm (Mundel et al., 1997). α-actinin-4, which is an actin cross-linker, directly interacts with Synpo via its C-terminus. This interaction allows Synpo to regulate actin bundling activity of α-actinin-4 by binding to both actin and α- actinin-4 (Asanuma et al., 2005). Synpo also regulates RhoA signaling and cell
240 Appendix 1 Synaptopodin project migration by protecting RhoA from proteosomal degradation, which promotes the stress fiber formation in podocytes (Asanuma et al., 2006).
Myosin 1e (Myo1e) belongs to Class I long tailed Myosin. It is broadly expressed in vertebrates with the highest expression pattern in lymph nodes, spleen and large intestine (Kim et al., 2006). In the kidney, it is enriched in podocytes, specialized epithelial cells. Myo1e plays an important role in podocytes function and normal glomerular filtration (Bi et al., 2013; Chase et al.,
2012). The SH3 domain of Myo1e, which is located at the C-terminal region of
Myo1e, was found to interact with dynamin and synaptojanin-1, playing a role in endocytosis (Krendel et al., 2007). Mundel lab discovered that Synaptopodin interacted with Myo1e likely through proline-SH3 domain interactions (private discussion with Dr. Peter Mundel). Therefore, we have decided to follow up on this possible interaction and investigate how it affects actin cytoskeleton in podocytes.
241 Appendix 1 Synaptopodin project
AP 1.2 Materials and Methods
PCR primers design
PCR primers were designed in a fashion that it includes 15 base pair (bp) extensions (5’) that are complementary to the ends of the linearized vector, which in this case is pFLAG-CMV-5a. Primer sequences are shown in AP Table 1.1.
In-Fusion® HD cloning
Cloning was done according to In-Fusion® HD cloning kit user manual from Clontech® (Mountain View, CA).
Western Blot analysis and Pulldown assay
Western blot analyses were done with anti-FLAG M2 (1:500) as primary antibody and HRP-GαM (1:3000) as secondary antibody.
Cos-7 cells were transiently transfected with desired plasmids. After 48 hours transfection, cells were lysed with ice-cold IP buffer (50 mM Tris-HCl pH7.6,
150 mM NaCl, 1% Triton X-100), followed by incubating on ice for 30 min. Cells then were centrifuged down in a table top microcentrifuge at 13,300 rpm at 4oC for 10 min. The supernatant (S1) was collected and the pellet was saved (P1). A small sample of the supernatant was used to prepare a gel sample. The rest of the supernatant was divided into three aliquots, and added to GST tagged proteins, followed by nutating at 4oC for 2 hours. At the end of two-hours nutation, a 50% slurry of BSA-coated glutathione-sepharose beads was added to each
242 Appendix 1 Synaptopodin project reaction, and the mixtures were nutated at 4 oC for 1 hour. Beads were centrifuged down at 4 oC at 500xg for 5 minutes and the supernatant was collected (Unbound). The beads were then washed with cold IP buffer for 5 times and then centrifuge down as above (Bound).
Immunofluorescence staining and Live-cell imaging
For immunofluorescence staining, mouse kidney podocytes were plated on collagen-IV coated coverslips and processed as previously described (Chase et al., 2012).
For live-cell imaging, wild type podocytes were plated and infected as previously described (Bi et al., 2013). 24 hours post infection, images of live cells were collected using a Nikon Eclipse TE2000-E multimode TIRF microscope equipped with a 60x, N.A. 1.45 TIRF Plan Apo Objective and a Hamamatsu EM-
CCD camera (Hamamatsu Photonics, Skukuza, Japan) and controlled by Nikon
Elements software.
Image manipulation
All fluorescence images were adjusted for brightness and contrast using
ImageJ software (National Institutes of Health, Bethesda, MA.)
243 Appendix 1 Synaptopodin project
AP 1.3 Results
AP1.3.1 Localization of Synaptopodin in podocytes
To examine the expression pattern of endogenous Synaptopodin in our podocytes clones, western blot analysis was performed on our Myo1e heterozygous and knock-out clones (AP Fig. 1.2A). The results show that
Synaptopodin is only expressed in differentiated Myo1e heterozygous and Myo1e knock-out podocytes clones, but not in any of the undifferentiated podocytes clones. This result is consistent with the fact that in the brain, Synaptopodin protein is only detected at a late stage of the development (Mundel and Kriz,
1995; Yamazaki et al., 2001). Using immunofluorescent staining, we were able to demonstrate the localization pattern of Synaptopodin in cultured differentiated podocytes (AP Fig. 1.2B), in which Synaptopodin decorates stress fibers (AP Fig.
1.2B, zoom-ins).
AP1.3.2 Generation and expression of FLAG-tagged Synaptopodin fragments
Synaptopodin is an actin-bundling protein and has multiple proline-rich regions, which makes it a rather linear shaped instead of globular shaped
(Mundel et al., 1997). Additionally, Proline-rich regions are thought to interact with SH3-domain containing proteins. To further map the interacting domains between Synaptopodin to Myo1e, PCR reaction and In-Fusion cloning methods were used to generate a few Synaptopodin fragments (Fig. 3A). Sequencing results and expression pattern in COS-7 cells revealed by western blot confirmed
244 Appendix 1 Synaptopodin project the accuracy of the cloning (Data not shown, and AP Fig. 1.3B). Not surprisingly, all the bands ran a little higher than expected size (AP Fig. 1.3B), likely due to the proline-rich regions in the constructs. Because of the low molecular weight of the last three short pieces of Synaptopodin fragments (Synpo-689-766aa, Synpo-
753-851aa, and Synpo-849-903aa), we were unable to visualize them on the western blot gel. Therefore, we used SynpoT (712-903 aa) fragment as a substitute for them (AP Fig. 1.1A and 1.3A).
AP1.3.3 Synaptopodin fragments interact with the Myo1e SH3 domain
To identify which Synaptopodin (Synpo) fragments specifically interacts with the Myo1e SH3 domain, we transfected Synpo 1-303aa, 299-504aa, 490-
700aa and Synpo-T into Cos-7 cells, and performed pulldown assays using GST- tagged purified proteins (GST, GST-Myo1e SH3, and GST-SH3WK) (AP Fig. 1.3C and D). All truncated Synpo proteins, as well as Synpo-Long, interacted with the
Myo1e SH3 domain in in vitro pull down assays (red boxes in AP Fig. 1.3C and
D). We also observed that some FLAG-tagged Synpo fragments precipitated with negative controls GST or GST-SH3wk. The possible explanation was when we removed the supernatant (unbound) from the pellet (beads, bound) it had residual supernatant containing FLAG-tagged Synpo fragments left with the pellet in the test tubes. Thus, using anti-FLAG antibody, the western blot detected
FLAG-tagged Synpo fragment bands under the negative control conditions.
However, this could be avoided by carefully separating supernatant from the pellet.
245 Appendix 1 Synaptopodin project
AP1.3.4 Synaptopodin does not co-localize with Myo1e in cultured podocytes
As previously investigated, Myo1e is enriched in cellular vesicles, cell-cell junctions, and plasma membranes (Bi et al., 2013; Krendel et al., 2007). Here, we want to demonstrate whether Synaptopodin colocalizes with Myo1e in any of these cellular structures. Cultured differentiated podocytes were infected with adenoviral vectors encoding mCherry-tagged Synaptopodin full length and GFP- tagged Myo1e full length. Live-cell images show no co-localization between
Synpo and Myo1e (AP Fig. 1.4). Since our in vitro pull down assays show that
Synaptopodin fragments specifically interact with the Myo1e-SH3 domain, we used GFP-tagged Myo1e-SH3 domain instead of the full-length protein. We introduced GFP-tagged Myo1e-SH3 domain into COS-7 cells via transient transfection. Surprisingly, no co-localizations were observed (data not shown).
Additionally, we used purified FLAG-tagged Synpo fragments to perform in vitro binding assay with GST-tagged Myo1e-SH3 domain, but the results all showed a consistent lack of binding (data not shown).
246 Appendix 1 Synaptopodin project
AP 1.4 Discussion
GST pull down assays indicated that Synpo interacted with the Myo1e
SH3 domain in vitro (AP Fig. 1.3C-D), with the strongest binding profiles of the N- terminal 1-303aa fragment and C-terminal Synpo-T fragment (712-903aa).
However, all microscopic data failed to confirm any Synpo-Myo1e SH3 binding interactions, since no significant co-localizations were observed during fluorescence microscopy analysis using either full-length Synpo and Myo1e (AP
Fig. 1.4), or only fragmented Synpo and the Myo1e-SH3 domain (data not shown). Synpo has been found to be an actin-associated protein that decorates the stress fibers in a dotted fashion, while Myosins belong to a family of actin- dependent molecular motors. Although Myo1e is not a processive motor, it has been confirmed that it has a weak binding affinity for actin in the presence of ATP
(El Mezgueldi et al., 2002). Because of the weak binding affinity for actin, Myo1e has been suggested to function in a region of high concentration of actin filaments (stress fibers), supposedly cross-linking them. Therefore, actin is potentially an ideal intermediate component to link Myo1e and Synpo together.
We hypothesize that the GST-tagged Myo1e-SH3 domain precipitates other proline-rich containing proteins that are directly associated with actin or Synpo.
To further investigate this matter, we used purified Synpo fragments to perform the in vitro binding assays to confirm whether this interaction is direct. However, the results showed consistent lack of binding. We have also performed co-IP experiments to prove the interaction, except our co-IP analysis failed to work due
247 Appendix 1 Synaptopodin project to unknown technical issues. An alterative method for investigating Synpo-Myo1e interaction is through mass spectrometry (Jonker et al., 2011; Kool et al., 2011), in which the identification of the Myo1e-SH3 domain binding partners could theoretically be deduce from podocytes lysate or tissue lysate after the precipitation by the Myo1e-SH3 domain.
Our previous investigation on mapping the truncated Myo1e using MDCK cells showed that GFP-tagged Myo1e-SH3 domain did not localize to any cellular structures, instead it was evenly distributed within the cytoplasm (Bi et al., 2013).
Therefore, using GFP-tagged Myo1e SH3 domain for microscopic analysis might not be a critical way to demonstrate the interaction between Synpo and Myo1e, since the SH3 domain itself might be lacking other significant binding motifs to function in vivo.
Our data has demonstrated lack of colocalization between Myo1e and
Synpo, even though they interact in vitro. This suggests that there might be a shared interacting partner by both Myo1e and Synpo. Further investigations could carry out to identify this protein.
248 Appendix 1 Synaptopodin project
AP 1.5 Reference:
Asanuma, K., Kim, K., Oh, J., Giardino, L., Chabanis, S., Faul, C., Reiser, J., Mundel, P., 2005. Synaptopodin regulates the actin-bundling activity of α-actinin in an isoform- specific manner. J. Clin. Invest. 115, 1188–98. doi:10.1172/JCI200523371.1188 Asanuma, K., Yanagida-Asanuma, E., Faul, C., Tomino, Y., Kim, K., Mundel, P., 2006. Synaptopodin orchestrates actin organization and cell motility via regulation of RhoA signalling. Nat. Cell Biol. 8, 485–91. doi:10.1038/ncb1400 Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M., 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am. J. Physiol. Renal Physiol. 305, 532–544. doi:10.1152/ajprenal.00223.2013 Chase, S.E., Encina, C. V, Stolzenburg, L.R., Tatum, A.H., Holzman, L.B., Krendel, M., 2012. Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am. J. Physiol. Renal Physiol. 303, F1099–106. doi:10.1152/ajprenal.00251.2012 El Mezgueldi, M., Tang, N., Rosenfeld, S.S., Ostap, E.M., 2002. The kinetic mechanism of Myo1e (human myosin-IC). J. Biol. Chem. 277, 21514–21. doi:10.1074/jbc.M200713200 Jonker, N., Kool, J., Irth, H., Niessen, W.M. A, 2011. Recent developments in protein- ligand affinity mass spectrometry. Anal. Bioanal. Chem. 399, 2669–2681. doi:10.1007/s00216- 010-4350-z Kim, S. V, Mehal, W.Z., Dong, X., Heinrich, V., Pypaert, M., Mellman, I., Dembo, M., Mooseker, M.S., Wu, D., Flavell, R. A, 2006. Modulation of cell adhesion and motility in the immune system by Myo1f. Science 314, 136–9. doi:10.1126/science.1131920 Kool, J., Jonker, N., Irth, H., Niessen, W.M. A, 2011. Studying protein-protein affinity and immobilized ligand-protein affinity interactions using MS-based methods. Anal. Bioanal. Chem. 401, 1109–1125. doi:10.1007/s00216-011-5207-9 Krendel, M., Osterweil, E.K., Mooseker, M.S., 2007. Myosin 1E interacts with synaptojanin-1 and dynamin and is involved in endocytosis. FEBS Lett. 581, 644–50. doi:10.1016/j.febslet.2007.01.021 Mundel, P., Gilbert, P., Kriz, W., 1991. Podocytes in glomerulus of rat kidney express a characteristic 44 KD protein. J. Histochem. Cytochem. 39, 1047–1056. doi:10.1177/39.8.1856454 Mundel, P., Heid, H.W., Mundel, T.M., Krüger, M., Reiser, J., Kriz, W., 1997. Synaptopodin: An actin-associated protein in telencephalic dendrites and renal podocytes. J. Cell Biol. 139, 193–204. doi:10.1083/jcb.139.1.193 Mundel, P., Kriz, W., 1995. Structure and function of podocytes: an update. Anat. Embryol. (Berl). 192, 385–97. Yamazaki, M., Matsuo, R., Fukazawa, Y., Ozawa, F., Inokuchi, K., 2001. Regulated expression of an actin-associated protein , synaptopodin , during long-term potentiation. J. Neurochem. 79, 192–199.
249 Appendix 1 Synaptopodin project
AP Figure 1.1. Domain mappings of Synaptopodin (Synpo) and Myosin 1e
(Myo1e).
250 Appendix 1 Synaptopodin project
AP Figure 1.1. Domain mappings of Synaptopodin (Synpo) and Myosin 1e
(Myo1e). (A) Three existing Synpo isoforms: Synpo-Long (1-903 aa), C-terminal
Synpo-short (1-685 aa), and N-terminal Synpo-T (712-903 aa). The yellow- colored boxes indicate the predicted proline-rich region in the protein based on sequence analysis (Mundel et al., 1997). (B) Myo1e protein contains a C-terminal globular head domain, a neck “IQ” motif that binds to calmodulin, and the tail domain that is comprised of membrane binding TH1 domain, proline-rich TH2 domain, and Src-homology (SH3) domain, which is responsible for protein- protein interactions.
251 Appendix 1 Synaptopodin project
AP Figure 1.2. Expression and localization of Synaptopodin in mouse podocytes.
252 Appendix 1 Synaptopodin project
AP Figure 1.2. Expression and localization of Synaptopodin in mouse podocytes. (A) Western blot analysis showing the expression of endogenous
Synpo in the lysates from several podocyte cell lines using rabbit Anti-
Synaptopodin NT (1:1000). Synpo is only expressed in differentiated podocytes, but not in undifferentiated ones. (B) Immunofluorescent staining images showing the co-localization of actin (red) and Synpo (green) in differentiated podocytes.
The “zoom-in” images show that Synpo localizes as puncta-like patterns on the stress fibers.
253 Appendix 1 Synaptopodin project
AP Figure 1.3. Synaptopodin interacts with Myo1e in pull down assays.
254 Appendix 1 Synaptopodin project
AP Figure 1.3. Synaptopodin interacts with Myo1e in pull down assays. (A)
FLAG-tagged Synpo fragments were generated by InFusion cloning and PCR, and confirmed by DNA sequencing. (B) Western blot analysis showing the expression of FLAG-tagged Synpo fragments from COS-7 cells lysates after transfection. (C-D) Western blot analysis of glutathione-S-transferase (GST) pulldown reactions using mouse anti-FLAG M2 (1:1000) antibody. GST-tagged
Myo1e-SH3 domain (GST-SH3) precipitated all the FLAG-tagged Synpo fragments (indicated by the red boxes).
255 Appendix 1 Synaptopodin project
AP Figure 1.4. Synaptopodin does not co-localize with full-length Myo1e in differentiated podocytes.
256 Appendix 1 Synaptopodin project
AP Figure 1.4. Synaptopodin does not co-localize with full-length Myo1e in differentiated podocytes. mCherry-tagged Synaptopodin full-length (A) and
GFP-tagged Myo1e full-length (A’) were introduced into mouse wild type podocytes using adenoviral vectors. The merged image (A’’) indicates no or minimal co-localization between Synaptopodin and Myo1e in live-cell imaging condition.
257 Appendix 1 Synaptopodin project
AP Table 1.1. Synpo-Long Infusion primer design
Note: Yellow highlighted bases are In Fusion sequences. Red –colored bases are Kozak sequences and start codon. Numbers inside the prentices indicate the melting temperature (Tm: oC) of each primer.
Synpo_L_F_1-303aa
TTGCGGCCGCAGATCTCACCATGGAGGGGTACTCAGAGGAGGCT (59.4)
Synpo_L_R_1-303aa
TACCGGATCCGTCGACTCGCTGCCCCATCATCCTCCT (62.4)
Synpo_L_F_299-504aa
TTGCGGCCGCAGATCTCACCATGGGGCAGCGAAGCCCG (62.6)
Synpo_L_R_299-504aa
TACCGGATCCGTCGACCTCTGGCGTGTGGCTTGAAGAC (58.5)
Synpo_L_F_490-700aa
TTGCGGCCGCAGATCTCACCATGGAGAAATATGTCATCGAGTCTTCA (56.3)
Synpo_L_R_490-700aa
TACCGGATCCGTCGACGTCTAGGCTGCTGGGGGGC (59.4)
Synpo_L_F_689-766aa
258 Appendix 1 Synaptopodin project
TTGCGGCCGCAGATCTCACCATGCCGGGCGCGTCCCGG (69.5)
Synpo_L_R_689-766aa
TACCGGATCCGTCGACCCGGGCCGCATTGATGATGTT (63.3)
Synpo_L_F_753-851aa
TTGCGGCCGCAGATCTCACCATGCAGGCGCTTCTGGCG (59.5)
Synpo_L_R_753-851aa
TACCGGATCCGTCGACGTAGAGGAAGGGCCTGGGAGG (57.7)
Synpo_L_F_847-903aa
TTGCGGCCGCAGATCTCACCATGCCCTTCCTCTACCGCCGC (63.1)
Synpo_L_R_847-903aa
TACCGGATCCGTCGACGGTGGTGCAGGAGGCCTCG (60.0)
259 Appendix 2 Myo1e-N164A project
Appendix Chapter 2
Investigation of the effects of the Myo1e
rigor mutant N164A on the dynamics of the
actin cytoskeleton
Appendix chapter 2 is written by myself and edited by my colleagues for grammatical errors. This chapter is not yet ready for publication, since there are still a few more experiments to perform in order to finish the story.
260 Appendix 2 Myo1e-N164A project
AP 2.1 Introduction
Myosins are actin-dependent molecular motors. In the past two decades, researchers have found more than 25 classes of myosins in eukaryotes.
Typically, myosins consist of 3 functional domains: the N-terminal motor domain that binds to actin in an ATP-dependent manner and is highly conserved through evolution, one or more IQ motifs for light chain binding, and the C-terminal tail domain which is highly diverse even within a single class. To study myosin functions and activity, researchers have used Dictyostelium discoideum (de la
Roche and Côté, 2001) and yeast (Brown, 1997; Kim and Flavell, 2008) as model systems, because they are less complex than most of the vertebrates.
In Dictyostelium discoideum, a point mutation, N233A, in the switch I loop of the myosin II motor domain was found to bind actin more tightly without moving the actin filaments in in vitro motility assays compared to wild type myosin II (Shimada et al., 1997). The switch I loop is located adjacent to the nucleotide binding pocket and is thought to play a role in nucleotide exchange during the ATPase cycle (Kintses et al., 2007). The rigor mutation N233A blocks the nucleotide binding by causing a conformational change to the switch I region and closing the nucleotide binding pocket, such that the mutant myosin II stays bound to actin filaments. Based on the conservation of the motor domain, another group of scientists generated an equivalent rigor mutation, N160A, in one of the vertebrate class I myosins, Myo1b, to test whether the motor activity is required for Myo1b function in controlling F-actin foci distribution. This N160A
261 Appendix 2 Myo1e-N164A project mutant was also found to be unable to move the F-actin (generating force) in vitro
(Almeida et al., 2011), which confirmed that the amino acid change from asparagine to alanine in myosin motor switch I loop from the previous study indeed turned myosin into a rigor mutant.
We have identified the equivalent asparagine in the switch I loop of Myo1e motor domain via sequence alignment, and introduced the same alanine mutation by mutagenesis (Myo1e N164A). In this study, we will investigate the effects of the Myo1e N164A mutation on the actin cytoskeleton dynamics.
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AP 2.2 Material and Methods
Cell cultures, transfection and infection
COS-7 cells were maintained in complete Dulbecco's Modified Eagle's
Medium (DMEM + 10% fetal bovine serum (FBS) + 1x antibiotic mix (ABM)).
Cells were plated in MatTek dishes for live-cell imaging. Immortalized Myo1e-null podocytes were cultured as previously described (Mundel et al., 1997).
Transient transfections were performed using JetPei transfection reagent according to manufacturer instructions. The infection of differentiated podocytes was performed as previously described appendix chapter III (Bi et al., 2014).
Live-cell imaging
Live-cell imaging was preformed as previously described in Chapter 2 (Bi et al., 2013)
Calcium switch assays
Calcium switch experiments were preformed as previously described in
Chapter 2 (Bi et al., 2013).
Constructions of plasmids encoding GST-TH2 protein and purification of GST fusion protein
The subcloning of Myo1e-TH2 domain into pGEX-6P-1 multiple cloning site was performed using InFusion® HD cloning kit (Clothech, Moutain View, CA).
263 Appendix 2 Myo1e-N164A project
The primers were used for cloning are sense (5’-3’)
GGGGCCCCTGGGATCCCCCAAGAACTCCCGTCCT, and anti-sense (5’-3’)
GTCGACCCGGGAATTCATAGGCTTACAAAGCCTTGCACTG. The restriction sites used within the primers are BamHI and EcoRI, which were underlined. A new stop codon (highlighted in red) at the end of the TH2 domain was also created. DNA sequencing confirmed the GST-TH2 is in frame with the sequences of the Myo1e-TH2 domain.
The expression and purification of the GST-TH2 was performed as previously described with modifications due to the high non-solubility of the TH2 domain (Jung and Hammer III, 1994). Bacterial E. coli strain Rosetta™ (DE3)
(Novagen, Merck Millipore, Billerica, MA) was inoculated at a low concentration, and the GST-TH2 protein expression was induced at OD595 = 0.8 with 1 mM final concentration of IPTG. After incubation for 3.5 hours at 30 oC, the bacterial cells were pelleted at 6000 g for 15 min at 4 oC, and the pellet was resuspended in ice cold buffer A (50 mM Tris-HCl, pH 8.0; 300 mM NaCl, 5 mM EDTA-Na, pH 7.5;
10% sarkosyl; 10 mM BME) with complete protease inhibitor (Roche) and 1mM
PMSF, followed by incubating at 4 oC over night. The next day, the lysate was centrifuged in a Ti-45 rotor at 15k rpm for 30 min at 4 oC (Beckman Instruments,
Inc., Fullerton, CA), and the pellet was discarded. The supernatant then was diluted with ice cold buffer B (50 mM Tris-HCl, pH 8.0; 300 mM NaCl, 5 mM
EDTA-Na, pH 7.5; 10 mM BME; 3.3% Triton X-100; 33 mM CHAPS) and checked for any precipitants (If there were any, the lysate was clarified by
264 Appendix 2 Myo1e-N164A project centrifugation as above and the pellet discarded). The final supernatant was loaded onto a Glutathione-Sepharose 4B column (Sigma, St. Louis, MO). The column was washed extensively with 5 column volumes (CV) of GST lysis buffer
(50 mM Tris-HCl, pH 8.0; 150 mM NaCl; 5 mM EDTA-Na, pH 7.5), 5 CV of
TBSE-150 buffer (20 mM Tris-HCl, pH 8.0; 150 mM NaCl; 1 mM EDTA), 5 CV of
TBSE-150 + 0.2% Thesit, 5 CV of TBSE-150 buffer, 5 CV of TBSE-150 + 1 M
NaCl, and 5 CV of TBSE-150 buffer, sequentially. The GST-TH2 fusion protein was eluted with TBSE-150 buffer containing 10 mM Glutathione. The fractions of the fusion protein were monitored by Bradford assay (Bio-Rad, Berkeley, CA) and confirmed by SDS-PAGE. Finally, the fractions enriched with the fusion protein were dialyzed against GST lysis buffer for twice, and twice more with GST lysis buffer containing 10% glycerol for long term storage. The concentration of the GST-TH2 fusion protein was determined by the Bio-Rad protein assay using bovine serum albumin (BSA) as a standard.
Actin purification and actin cosedimentation assays
The rabbit skeletal muscle acetone powder was purchased from Pelfreez
(Pelfreez-bio, Rogens, AR). The purification of rabbit skeletal muscle actin was preformed as previously described (Pardee and Spudich, 1982) with minor modifications. Briefly, the acetone powder was mixed with ice cold buffer G (2 mM Tris-HCl, pH 8.0; 0.2 mM ATP; 0.5 mM DTT; 0.2 mM CaCl2) and stirred at 4 oC for 30 min. The extract was then centrifuged at 35K rpm (142K g) for 15 min at
4 0C in Beckman 70-Ti ultracentrifuge rotor (Beckman Coulter, Pasadena, CA).
265 Appendix 2 Myo1e-N164A project
Following the centrifugation, the volume of the supernatant was measured using a graduated cylinder. Then the supernatant was stirred in a beaker at room temperature while adding KCl, MgCl2 and ATP to final concentrations of 50 mM,
2 mM, and 1 mM, respectively, as well as protease inhibitor cocktails (Pefabloc,
Roche), followed by incubating at room temperature without stirring for 30 min.
Then the mixture was transferred to 4 oC and incubated for 60 min without stirring.
At the end of the incubation, KCl was added to the mixture to a final concentration of 0.8 M while stirring. After the addition of KCl, the mixture was stirred at 4 oC for additional 60 min before being centrifuged in a Beckman 70-Ti rotor at 35K rpm for 3 hours. At the end of the centrifugation, the supernatant was removed and the pellet was washed with ice cold buffer G gently 5 times to remove the excess salt while not to dislodging the pellet. The pellet (F-actin) was then scraped out using a Teflon spatula and resuspended in ice cold buffer G using a loose glass pestle in a Kontes homogenizer. The resuspended G-actin solution was then dialyzed 3 times against fresh buffer G, and clarified by centrifugation in Beckman 70-Ti rotor at 35K rpm for 3 hours. The concentration of clarified G-actin solution was determined by absorbance at 280 nm, and the molecular weight was confirmed by SDS-PAGE.
The actin cosedimentation assays were performed according to previously described procedure (Goode et al., 1998). Purified G-actin was diluted to a proper stock concentration (in this case, we used 20 μM) in buffer G and polymerized in F-buffer (G-buffer + 1x Initiation mix) for 1 hour at room
266 Appendix 2 Myo1e-N164A project temperature before using in the actin co-sedimentation assay. At this point, 20
μM polymerized actin was a mixture of 0.2 μM actin monomer and 19.8 μM actin polymer. Then, 2 μM freshly polymerized actin was mixed with
0/0.2/0.4/0.8/1/2/4/8/16/20 μM purified GST-TH2 fusion protein (for a total volume of 50 μL) and incubated at room temperature for 10 min. The mixtures were sedimented by centrifugation at 90,000 rpm for 20 min at 20 oC in a TLA-
100 rotor (Beckman Coulter, Pasadena, CA). The supernatants and the pellets were separated and the pellets were resuspended in equal amount of 1x sample buffer. Equal amounts of the supernatants and pellets suspensions analyzed by
SDS-PAGE. The gels were Coomassie-stained and the intensity of the GST-TH2 and actin in the pellet was quantified using ImageJ software (National Institute of
Health, Bethesda, Maryland). The data was plotted and fitted in KaleidaGraph
(Synergy Software) using the following equation that was derived from (Sirotkin et al., 2005):
Y=((M0+2+M1)-((M0+2+M1)2-4*M0*2)0.5)/4; M1=0.1
Y is the fraction of ligand bound; M0 is [GST-TH2] at variable concentrations; 2 refers to the soluble ligand at a constant concentration (In this case, [actin]=2
μM); M1 is the dissociation constant Kd.
Quantification of the protrusions
We categorized the protrusions based on their length and width. Regular filopodia have a length of less than 1.5 μm and are spike-like (no width).
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Enlongated protrusions have a length between 1.5 to 12 μm, and a width less than line width of 2 when set up parameter in ImageJ. The bulbed protrusions have lengths longer than 12 μm, and the width of the protrusions were greater than line width of 2 when set up parameter in ImageJ. A total of 146 protrusions in wild type Myo1e expressing cells and 66 protrusions in Myo1e-N164A expressing cells were measured. All the measurements were done in ImageJ
(National Institute of Health, Bethesda, MD), and the data were plotted as bar graphs in KaleidaGraph (Synergy Software).
Fluorescence recovery after photobleaching (FRAP)
The FRAP experiments were preformed as described in Material and
Methods in Chapter 3.
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AP 2.3 Results
AP 2.3.1 The rigor mutant Myo1e N164A co-localizes with actin at cell protrusions and plasma membranes.
Wild type Myo1e localizes to plasma membranes, cell-cell junctions and endocytic vesicles. To investigate the localization pattern of the rigor mutant
N164A relative to actin, we transiently transfected COS-7 cells with mCherry- tagged Lifeact and GFP-tagged Myo1e-N164A (AP Fig. 2.1C-D’’). In cells expressing the N164A mutant, we noticed that there appeared to be longer actin- rich protrusions forming with N164A decorating along the protrusions (AP Fig.
2.1C-C’’, yellow arrows). Additionally, the plasma membrane localization of the
N164A mutant was much more prominent (AP Fig. 2.1D’, green arrows) compared to the wild type Myo1e expressing cells (AP Fig. 2.1A’-B’). In both wild type Myo1e and the Myo1e-N164A expressing cells, we found the formation of the regular “spike-like” filopodia (AP Fig. 2.1B, red arrows). From these results we have concluded that the rigor mutant Myo1e N164A has prominent membrane localizations and induces the formation of long protrusions in COS-7 cells. However, the nature of these protrusions is unknown, and still under investigation.
We categorized the protrusions in COS-7 cells expressing wild type
Myo1e or the rigor mutant Myo1e N164A (AP Fig. 2.2A). Regular filopodia are the normal actin-rich protrusions, which are found in most of the cells. These are
269 Appendix 2 Myo1e-N164A project characterized by the protrusions being shorter than 1.5 μm and spike-like (AP Fig.
2.2A, upper panel). The elongated protrusions are characterized by having lengths between 1.5 to 12 μm. The bulbous protrusions are characterized as protrusions whose lengths are greater than 12 μm, and decorated by myosin puncta (bulb-shaped protrusions). Bulbous protrusions mostly have a width greater than 2 when measured in ImageJ. We found that cells expressing wild type Myo1e form more regular protrusions, while almost 80% of cells expression the rigor mutant are bulb-shaped (AP Fig. 2.2B).
AP 2.3.2 The rigor mutant Myo1e N164A co-localizes with ZO-1 at cell-cell junctions.
To examine whether the Myo1e N164A mutant affects ZO-1 localization,
Myo1e null mouse podocytes were infected with adenoviral vectors encoding mCherry-tagged ZO-1 (AP Fig. 2.3A-C) and GFP-tagged Myo1e wild type (AP
Fig. 2.3A’-B’) or N164A mutant (AP Fig. 2.3C’). The rigor mutant Myo1e N164A co-localized with ZO-1 at cell-cell junctions, similar to Myo1e wild type (AP Fig.
2.3, yellow arrows). Interestingly, the Myo1e N164A mutant was enriched in the plasma membrane, which was also observed in COS-7 cells (AP Fig. 2.3C’-C’’, green arrows).
AP 2.3.3 The rigor mutant Myo1e N164A is recruited to nascent cell-cell contacts with ZO-1 during calcium switch experiments.
270 Appendix 2 Myo1e-N164A project
To further study the effect of N164A mutant on ZO-1 activity, we performed calcium switch experiments as previously described in Chapter 2 (Bi et al., 2013) (AP Fig. 2.4). As expected, the Myo1e N164A mutant was recruited to nascent cell-cell contacts during junction assembly, along with ZO-1 (AP Fig.
2.4, yellow arrows).
AP 2.3.4 The rigor mutant Myo1e N164A does not alter ZO-1 dynamics at the junctions.
To investigate whether the Myo1e N164A mutant affects ZO-1 dynamics at the junctions, we performed fluorescent recovery after photobleaching (FRAP) experiments on Myo1e null mouse podocytes that were re-introduced with either wild type Myo1e or the Myo1e N164A mutant (AP Fig. 2.5). The data was fit to a double exponential curve (AP Fig 2.5A), and the mobile fractions were calculated and displayed in AP Fig. 2.5B. The junctional ZO-1 fluorescence recovery was
70%, similar to that in the cells expressing wild type Myo1e. This shows that the
Myo1e-N164A mutant does not alter ZO-1 dynamics at the junctions.
AP 2.3.5 Deletion of the TH2 domain from the Myo1e N164A mutant mimics the phenotype of Myo1e-ΔTH2 in the cells.
As evidenced in Chapter 2, the TH2 domain of Myo1e is essential for targeting Myo1e to cell-cell contacts. Since the N164A mutant is supposed to be a rigor mutant, we hypothesized that introducing a rigor mutation to the motor domain will rescue the Myo1e-ΔTH2 localization in the cells from diffuse to the
271 Appendix 2 Myo1e-N164A project junctions. We generated Myo1e-N164A-ΔTH2 by mutagenesis and transfected it into COS-7 cells, along with mCherry-tagged Lifeact (AP Fig. 2.6). By comparing and analyzing the distributions of Myo1e-ΔTH2 (AP Fig. 2.6A-A’’ and 2.6C-C’’) and Myo1e-N164A-ΔTH2 (AP Fig. 2.6B-B’’ and 2.6D-D’’) in both single cells (AP
Fig. 2.6A-B’’) and cell pairs (AP Fig. 2.6C-D’’), we could not find any significant differences in localization between Myo1e-ΔTH2 and Myo1e-N164A-ΔTH2, suggesting that TH2 domain is indeed essential for Myo1e localization in the cell.
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AP 2.4 Discussion
AP 2.4.1 Is Myo1e N164A a rigor mutant?
We have predicted that the Myo1e N164A mutant is a rigor binding mutant based on previous studies (Almeida et al., 2011; Shimada et al., 1997). We attempted to confirm this prediction by live-cell imaging. Here we have shown that the localization patterns of Myo1e N164A in cells are very prominent at the plasma membrane and at cell-cell junctions. As we know, the underlying cortical actin filaments plays a key role in maintaining plasma membrane shape and generating tensile force for initiating proper cell processes such as migration and endocytosis. According to previous studies, due to its low affinity to actin filaments, Myo1e has been suggested to be enriched in places that have a high concentration of short actin filaments (El Mezgueldi et al., 2002). Additionally, our lab, among other groups, have demonstrated that there is a second actin-binding site at Myo1e tail domain (AP Fig. 2.7, and Brzeska et al., 1988; Doberstein and
Pollard, 1992; Jung and Hammer III, 1994; Lynch et al., 1986; Swanljung-Collins and Collins, 1994; Lee et al., 1999; Rosenfeld and Rener, 1994). The affinity of
Myo1e to actin ranges from 0.1 μM to 7.2 μM depending on salt and ATP concentrations. Our data suggested that the tail domain has a second actin- binding site with the affinity to actin of 3.6 μM. These two weak actin-binding sites, one in the motor domain and the other in the tail domain, might contribute
Myo1e localization to actin-rich regions in the cells and its role in cross linking short actin filaments underlying the plasma membrane. Therefore, we have
273 Appendix 2 Myo1e-N164A project hypothesized that the N164A mutant increases Myo1e binding affinity to actin, which could account for the prominent localization to plasma membranes.
However, further in vitro motility assays are necessary to confirm this prediction.
In the live cell images of podocytes expressing N164A, we also observed more bright vesicles in the cytoplasm, which lead us to the hypothesis that the
N164A mutant might promote endocytosis. Since this mutant is predicted to be a rigor mutant, theoretically it should bind to actin filaments tighter and spend more time on actin filaments compared to wild type Myo1e. We attempted to confirm this by immunostaining other proteins that are involved in the endocytic pathways, such as Dynamin-2, Rab11, EEA or Clathrin (data not shown). We hypothesized that if the rigor mutant promotes endocytosis by binding to the actin filament tighter, we would observe an increasing co-localization pattern with the endocytic proteins in Myo1e-N164A expressing cells. Unfortunately, we failed to detect hypothesized phenomenon (data not shown). Therefore, it suggests that the
N164A positive vesicles we saw in the cells are unlikely to be involved in endocytosis.
AP 2.4.2 What is the nature of the long protrusions that are induced by the
Myo1e N164A mutant?
The Myo1e N164A mutant induces the formation of long bulbed protrusions in both COS-7 cells (AP Fig. 2.1) and podocytes (data not shown).
Live cell imaging with mCherry-tagged Lifeact indicated that these long bulbed
274 Appendix 2 Myo1e-N164A project protrusions are rich in actin. We have hypothesized that by increasing the affinity to actin filament near the plasma membrane, the Myo1e N164A mutant promotes actin polymerization by stabilizing actin. Our preliminary imaging data with mCherry-tagged EMTB (the microtubule binding domain of ensconsin) suggested that these long bulb-like protrusions are also rich in microtubule (data not shown).
This suggests that the nature of the long protrusions we saw in cells expressing the Myo1e N164A mutant is different from the novel filopodia. The microtubules based long protrusions have been found in neuronal cells during the induction of axons (Goldberg and Burmeister, 1992). In the future investigations, we may further explore the nature of Myo1e N164A induced long protrusions by examining the pathways that promote the formation of long filopodia in cells.
AP 2.4.3 What is the effect of the rigor mutant on the dynamics of the actin cytoskeleton?
Class I myosins are predicted to be involved in the regulation of membrane tension (Nambiar et al., 2009). Recent preliminary studies from our lab have also shown that the membrane tension is reduced 25% in Myo1e/Myo1f double knockout microphages (unpublished data-S.B.). If the Myo1e N164A mutant is a true rigor mutant, then it should result in increased membrane tension.
According to the model that Lieber et al. proposed, actin polymerization creates a local force at the cell leading edge, and the cell counterbalances this force through generating membrane tension (Lieber et al., 2013). We propose that the
Myo1e N164A stabilizes the actin filament by staying bound to it, allowing the
275 Appendix 2 Myo1e-N164A project actin to continue to polymerize. The actin will push against the membrane more, thus increasing the membrane tension. Actin polymerization at the leading edge also promotes lamellipodia and filopodia formation and in turn regulates cell migration (Chen et al., 2014; Pollard et al., 2000; Vicente-Manzanares et al.,
2005). Therefore, we would hypothesize that the Myo1e N164A mutant would also affect cell migration. This also correlates our data that the N164A mutant induces the formation of long protrusions, which is the initiation step of cell migration.
AP 2.4.4 Is motor activity required for Myo1e function at the junctions?
Myo1e, as one of the Class I myosins, has been characterized as a low duty ratio motor, which means it only spends a short amount of time bound to actin, and this process is coupled with the release of the inorganic phosphate (De
La Cruz and Ostap, 2004; El Mezgueldi et al., 2002). Based on previous studies, we predicted that introducing the N164A point mutation to the Myo1e motor domain would create a rigor binding and motor dead mutant. This means the
Myo1e N164A should stay bound to actin filaments in an ATP-independent manner, but would not be able to generate force. As an actin-binding myosin without motor activity, we can explore whether the actin binding ability, or the motor activity, or both are required for Myo1e functions at the junctions. Our
FRAP analysis showed no differences in ZO-1 exchange at the junctions in wild type Myo1e and the Myo1e N164A expressing cells. This indicated that the motor activity of Myo1e is not required for regulating ZO-1 dynamics at the junctions,
276 Appendix 2 Myo1e-N164A project but only the actin binding ability. A follow-up analysis of FRAP experiments on junctional wild type Myo1e and the Myo1e-N164A mutant would allow us to elucidate whether there are any differences in Myo1e exchange at the junctions without Myo1e motor activity.
In conclusion, we have shown that the Myo1e N164A mutant prominently localizes to the plasma membrane and intracellular vesicles, and colocalizes with actin and ZO-1 at cell-cell contacts, but there is not enough evidence to confirm its function and activity in cells. Therefore, further detailed biochemical characterizations are highly recommended.
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AP 2.5 Reference:
Almeida, C.G., Yamada, A., Tenza, D., Louvard, D., Raposo, G., Coudrier, E., 2011. Myosin 1b promotes the formation of post-Golgi carriers by regulating actin assembly and membrane remodelling at the trans-Golgi network. Nat. Cell Biol. 13, 779–789. doi:10.1038/ncb2262 Bi, J., Chase, S.E., Pellenz, C.D., Kurihara, H., Fanning, A.S., Krendel, M., 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am. J. Physiol. Renal Physiol. 305, 532–544. doi:10.1152/ajprenal.00223.2013 Bi, J., Pellenz, C.D., Krendel, M., 2014. Visualization of cytoskeletal dynamics in podocytes using adenoviral vectors. Cytoskeleton (Hoboken). 71, 145–156. doi:10.1016/j.biotechadv.2011.08.021.Secreted Brown, S.S., 1997. Myosins in yeast. Curr. Opin. Cell Biol. 9, 44–8. Brzeska, H., Lynch, T.J., Korn, E.D., 1988. Localization of the actin-binding sites of Acanthamoeba myosin IB and effect of limited proteolysis on its actin-activated Mg2+-ATPase activity. J. Biol. Chem. 263, 427–35. Chen, C., Tao, T., Wen, C., He, W. Q., Qiao, Y. N., Gao, Y. Q., Chen, X., Wang, P., Chen, C. P., Zhao, W., Chen, H. Q., Ye, A. P., Peng, Y. J., Zhu, M. S., 2014. Myosin Light Chain Kinase (MLCK) Regulates Cell Migration in a Myosin Regulatory Light Chain Phosphorylation-Independent Mechanism. J. Biol. Chem. 289, 0–23. doi:10.1074/jbc.M114.567446 De La Cruz, E.M., Ostap, E.M., 2004. Relating biochemistry and function in the myosin superfamily. Curr. Opin. Cell Biol. 16, 61–7. doi:10.1016/j.ceb.2003.11.011 De la Roche, M. a, Côté, G.P., 2001. Regulation of Dictyostelium myosin I and II. Biochim. Biophys. Acta 1525, 245–61. Doberstein, S.K., Pollard, T.D., 1992. Localization and specificity of the phospholipid and actin binding sites on the tail of Acanthamoeba myosin IC. J. Cell Biol. 117, 1241–9. El Mezgueldi, M., Tang, N., Rosenfeld, S.S., Ostap, E.M., 2002. The kinetic mechanism of Myo1e (human myosin-IC). J. Biol. Chem. 277, 21514–21. doi:10.1074/jbc.M200713200 Goldberg, D.J., Burmeister, D.W., 1992. Microtubule-based filopodium-like protrusions form after axotomy. J. Neurosci. 12, 4800–4807. Goode, B.L., Drubin, D.G., Lappalainen, P., 1998. Regulation of the cortical actin cytoskeleton in budding yeast by twinfilin, a ubiquitous actin monomer-sequestering protein. J. Cell Biol. 142, 723–733. doi:10.1083/jcb.142.3.723 Jung, G., Hammer III, J.A., 1994. The actin binding site in the tail domain of Dictyostelium myosin IC ( tail homology region 2 ). FEBS Lett. 342, 197–202. Kim, S. V, Flavell, R. A, 2008. Myosin I: from yeast to human. Cell. Mol. Life Sci. 65, 2128–37. doi:10.1007/s00018-008-7435-5 Kintses, B., Gyimesi, M., Pearson, D.S., Geeves, M. A, Zeng, W., Bagshaw, C.R., Málnási-Csizmadia, A., 2007. Reversible movement of switch 1 loop of myosin determines actin interaction. EMBO J. 26, 265–274. doi:10.1038/sj.emboj.7601482 Lee, W. L., Ostap, E.M., Zot, H.G., Pollard, T.D., 1999. Organization and Ligand Binding Properties of the Tail of Acanthamoeba Myosin-IA. (IDENTIFICATION OF AN ACTIN- BINDING SITE IN THE BASIC (TAIL HOMOLOGY-1) DOMAIN). J. Biol. Chem. 274, 35159– 35171. doi:10.1074/jbc.274.49.35159 Lieber, A.D., Yehudai-Resheff, S., Barnhart, E.L., Theriot, J. A., Keren, K., 2013. Membrane tension in rapidly moving cells is determined by cytoskeletal forces. Curr. Biol. 23, 1409–1417. doi:10.1016/j.cub.2013.05.063
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Lynch, T.J., Albanesi, J.P., Korn, E.D., Robinson, E. A, Bowers, B., Fujisaki, H., 1986. ATPase activities and actin-binding properties of subfragments of Acanthamoeba myosin IA. J. Biol. Chem. 261, 17156–62. Mundel, P., Reiser, J., Kriz, W., 1997. Induction Podocytes of Differentiation in Cultured Rat and Human. Am. Soc. Nephrol. 8, 697–705. Nambiar, R., McConnell, R.E., Tyska, M.J., 2009. Control of cell membrane tension by myosin-I. Proc. Natl. Acad. Sci. U. S. A. 106, 11972–11977. doi:10.1073/pnas.0901641106 Pardee, J.D., Spudich, J.A., 1982. Purification of muscle actin. Methods Enzymol. 85, 164–181. Pollard, T.D., Blanchoin, L., Mullins, R.D., 2000. Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu. Rev. Biophys. Biomol. Struct. 29, 545–76. Rosenfeld, S.S., Rener, B., 1994. The GPQ-rich segment of Dictyostelium myosin IB contains an actin binding site. Biochemistry 33, 2322–28. Shimada, T., Sasaki, N., Ohkura, R., Sutoh, K., 1997. Alanine scanning mutagenesis of the switch I region in the ATPase site of Dictyostelium discoideum myosin II. Biochemistry 36, 14037–14043. doi:10.1021/bi971837i Sirotkin, V., Beltzner, C.C., Marchand, J.-B., Pollard, T.D., 2005. Interactions of WASp, myosin-I, and verprolin with Arp2/3 complex during actin patch assembly in fission yeast. J. Cell Biol. 170, 637–48. doi:10.1083/jcb.200502053 Swanljung-Collins, H., Collins, J.H., 1994. Brush border myosin I has a calmodulin/phosphatidylserine switch and tail actin-binding.pdf, Actin: Biophysics, Biochemistry, and Cell Biology. Vicente-Manzanares, M., Webb, D.J., Horwitz, A.R., 2005. Cell migration at a glance. J. Cell Sci. 118, 4917–4919. doi:10.1242/jcs.02662
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AP Figure 2.1. The Myo1e N164A mutant co-localizes with actin at the plasma membranes and cell protrusions.
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AP Figure 2.1. The Myo1e N164A mutant co-localizes with actin at the plasma membranes and cell protrusions. COS-7 cells were transiently transfected with mCherry-tagged Lifeact (A-D) and GFP-tagged wild type Myo1e
(A’-B’) or the Myo1e N164A mutant (C-D’). The yellow arrows indicate the co- localization of the Myo1e N164A mutant and actin at the cell protrusions. The green arrows (D’) demonstrate the prominent plasma membrane enrichment of the Myo1e-N164A mutant. The red arrows (B) point out the formation of the regular filopodia. Scale bar: 10 μm.
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AP Figure 2.2. Categories and quantifications of the protrusions in COS-7 cells.
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AP Figure 2.2. Categories and quantifications of the protrusions in COS-7 cells. (A) Representative examples of the three categories of protrusions found in COS-7 cells expressing wild type Myo1e or the Myo1e-N164A mutant. (B) The percentage of each category is presented in bar graph.
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AP Figure 2.3. The Myo1e N164A mutant co-localizes with ZO-1 at the cell- cell junctions, similar as the wild type Myo1e.
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AP Figure 2.3. The Myo1e N164A mutant co-localizes with ZO-1 at the cell- cell junctions, similar as the wild type Myo1e. Myo1e-null mouse podocytes were infected with adenoviral vectors encoding mCherry-tagged ZO-1 (A-C) and
GFP-tagged wild type Myo1e (A’-B’) or the Myo1e-N164A mutant (C’). Yellow arrows indicate the co-localization of ZO-1 and Myo1e at the cell-cell junctions, while the green arrows point out the prominent plasma membrane localization of the Myo1e-N164A mutant (C’-C’’). Scale bar: 10 μm.
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AP Figure 2.4. The Myo1e N164A mutant is recruited to the nascent cell-cell contact with ZO-1 during contact assembly.
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AP Figure 2.4. The Myo1e N164A mutant is recruited to the nascent cell-cell contact with ZO-1 during contact assembly. Myo1e-null mouse podocytes were infected with adenoviral vectors encoding GFP-tagged Myo1e-N164A mutant and mCherry-tagged ZO-1. The videos were taken during calcium switch assays to mimic the junction assembly. Representative frames from the videos were taken at 5 min, 10 min and 15 min time point. The yellow arrows in the right panels indicate the recruitment of both Myo1e N164A mutant and ZO-1 during nascent contact formation. Scale bar: 10 μm.
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AP Figure 2.5. The Myo1e N164A mutant does not alter the dynamics of junctional ZO-1.
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AP Figure 2.5. The Myo1e N164A mutant does not alter the dynamics of junctional ZO-1. Fluorescence recovery after photobleach (FRAP) experiments were performed in Myo1e-null mouse podocytes re-expressing wild type Myo1e or the Myo1e-N164A mutant. (A) The recovery of junctional ZO-1 intensity was recorded and calculated, and fitted into a double exponential curve. The error bars indicate standard deviations. (B) The mobile fractions of mCherry-tagged
ZO-1 plotted using KaleidaGraph (Synergy Software).
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AP Figure 2.6. N164A mutation does not rescue cellular distribution of
Myo1e-ΔTH2.
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AP Figure 2.6. N164A mutation does not rescue cellular distribution of
Myo1e-ΔTH2. COS-7 cells were co-transfected with GFP-tagged Myo1e-ΔTH2
(A and C) or Myo1e-N164A-ΔTH2 (B and D), and mCherry-tagged Lifeact (A’-D’).
The images contain single cells are depicted in A-B’’, while the cell pairs are in C-
D’’. Scale bar: 10 μm.
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AP Figure 2.7. Actin cosedimentation assay shows the binding curve of 2
μM GST-TH2 fusion protein and increasing amount of F-actin.
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AP Figure 2.7. Actin cosedimentation assay shows the binding curve of 2
μM GST-TH2 fusion protein and increasing amount of F-actin. The representative curve was generated and fitted in KaleidaGraph (Synergy
Software). The Kd of GST-TH2 is 3.6 μM.
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Appendix Chapter 3
Visualization of cytoskeletal dynamics in
podocytes using adenoviral vectors
Appendix chapter 3 is adapted from the following article published in 2014: Bi, J., Pellenz, C.D., Krendel, M., 2014. Visualization of cytoskeletal dynamics in podocytes using adenoviral vectors. Cytoskeleton (Hoboken). 71, 145–156.
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AP 3.1 Attributions and Acknowledgments
Pellenz, CD and I are co-first authors on this publication. Pellenz, CD designed all viral production procedures with modifications from the manufecture suggestions. I performed all the experiments with the help from Pellenz, CD, and prepared all the figures. Pellenz, CD, wrote the materials and methods session, while I finished writing the rest of the draft. Dr. Krendel, M edited the manuscipt for publication.
The authors would like to thank Peter Mundel for generously sharing podocyte cell line and synaptopodin constructs and for his expert advice on podocyte biology and cell culture techniques. The authors are also grateful for help and advice on working with podocytes provided by Christian Faul, Kirk
Campbell, and Michelle Rheault. The authors thank Alan Fanning for providing
ZO-1 constructs and Pietro De Camilli and Roberto Zoncu for sharing clathrin light chain construct.
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AP 3.2 Abstract
Glomerular visceral epithelial cells (podocytes) play a key role in maintaining selective protein filtration in the kidney. Podocytes have a complex cell shape characterized by the presence of numerous actin-rich processes, which cover the surface of glomerular capillaries and are connected by specialized cell-cell adhesion complexes (slit diaphragms). Human genetic studies and experiments in knockout mouse models show that actin filaments and actin-associated proteins are indispensable for the maintenance of podocyte shape, slit diaphragm integrity, and normal glomerular filtration. The ability to examine cytoskeletal protein organization and dynamics in podocytes and to test the effects of disease-associated mutations on protein localization provides valuable information for researchers aiming to dissect the molecular mechanisms of podocyte dysfunction. We describe how adenovirus-mediated transduction of cultured podocytes with DNA constructs can be used to reliably introduce fluorescently tagged cytoskeletal markers for live cell imaging with high efficiency and low toxicity. This technique can be used to study the dynamic reorganization of the podocyte cytoskeleton and to test the effects of novel mutations on podocyte cytoskeletal dynamics.
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AP 3.3 Introduction
Renal glomeruli play an important role in selective filtration in the kidney by allowing the passage of water, small solutes, and waste products into the primary urinary filtrate while retaining high molecular weight proteins. Glomerular visceral epithelial cells, also known as podocytes, are essential for selective protein filtration in the glomerulus [Greka and Mundel 2012; Pavenstadt et al.
2003].
Podocytes extend numerous interdigitating processes (foot processes), which cover the entire surface of glomerular capillaries and are interconnected by specialized cell-cell junctions, known as slit diaphragms. Loss of podocytes or disruption of the podocyte foot process architecture and slit diaphragm integrity results in defects in protein filtration, leading to proteinuria (protein in the urine)
[Greka and Mundel 2012; Reiser and Sever 2013]. Nephrotic syndrome, a disease state characterized by the massive proteinuria due to the loss of selective glomerular filtration, is a serious health-threatening condition, which can lead to chronic kidney disease and renal failure. Disruption of glomerular filtration significantly increases the risk of death and cardiovascular complications in affected patients. Therefore, understanding molecular mechanisms leading to glomerular dysfunction is of major health importance.
Podocyte cytoskeleton is necessary for the maintenance of normal podocyte structure, and disruption of the cytoskeletal organization in podocytes leads to the flattening (effacement) of foot processes, loss of slit diaphragm
297 Appendix 3 Technique paper integrity, and defects in filtration [Faul et al. 2007; Mathieson 2012; Welsh and
Saleem 2012]. Human genetic studies have identified mutations in several components of the actin cytoskeleton that are associated with glomerular dysfunction. These include actin crosslinker α-actinin-4, actin assembly regulator
INF2, scaffolding protein CD2AP, and actin-dependent molecular motor myosin
1e [Brown et al. 2010; Kaplan et al. 2000; Kim et al. 2003; Mele et al. 2011;
Sanna-Cherchi et al. 2011]. Many glomerular disorders have also been linked to the proteolytic degradation of the important regulators of podocyte actin organization, for example, synaptopodin [Mundel and Reiser 2010], further underscoring the importance of the actin cytoskeleton for normal podocyte functions. The ability to characterize the intracellular localization of normal and mutant cytoskeletal proteins and the effects of disease-associated mutations on the cytoskeletal dynamics in podocytes is important in order to establish the mechanistic links between specific mutations found in patients and the corresponding glomerular defects.
Many studies of podocyte biology that require detailed characterization of molecular mechanisms underlying glomerular disease rely on the use of cultured podocytes. Primary podocyte cultures can be derived from explants growing out from isolated glomeruli, which are obtained using differential sieving of the kidney cortex or magnetic bead isolation and plated onto collagen-coated culture dishes
[Katsuya et al. 2006; Shankland et al. 2007]. Derivation of podocyte explants from the transgenic mouse strain carrying a temperature-sensitive, interferon-
298 Appendix 3 Technique paper inducible version of the immortalizing large T-antigen of SV-40 virus
(Immortomouse) [Jat et al. 1991], allows establishment of stable, conditionally immortalized podocyte cell lines [Mundel et al. 1997b; Schiwek et al. 2004].
Alternatively, podocyte cell lines (for example, those derived from human glomeruli) can be conditionally immortalized by the retrovirus-mediated expression of the temperature-sensitive large T antigen [Ni et al. 2012; Saleem et al. 2002]. Conditionally immortalized podocytes can be maintained under the permissive conditions (33°C + interferon) and switched to non-permissive conditions (37°C or 38°C) in order to induce differentiation. A detailed discussion of the technical challenges and advantages of maintaining podocytes in culture is provided in [Shankland et al. 2007]. While differentiated podocytes in culture do not fully recapitulate the complex architecture of podocytes in vivo, they do express many proteins characteristic of podocytes and represent a valuable in vitro model for testing the effects of disease-associated mutations. An additional advantage of conditionally immortalized podocyte lines is the ability to derive these cells from knockout animal models lacking the protein of interest. These knockout podocytes can then be used to reintroduce the wild type or mutant protein and to examine its effects on the podocyte phenotype [Bi et al. 2013].
One of the challenges in working with conditionally immortalized podocytes is the low efficiency of expression of recombinant proteins (for example, fluorescently tagged proteins for live imaging studies). Non- differentiated podocytes can be transiently transfected using standard
299 Appendix 3 Technique paper transfection reagents; however, the transfection efficiency, even under optimized conditions, rarely exceeds 20% [Shankland et al. 2007]. Since a typical experiment utilizing differentiated podocytes requires 7-14 days of growth under non-permissive conditions to obtain fully differentiated cells, plasmids introduced by transient transfection are likely to be lost. Alternatively, stably transfected cell lines can be derived using either a transient transfection or lentiviral infection of non-differentiated podocytes with subsequent antibiotic selection [Kaufman et al.
2007; Reiser et al. 2004]. Establishment of stable cell lines is a lengthy process and it limits the ability to introduce multiple markers into the same cell line (for example, two proteins with distinct fluorescence tags).
Live cell imaging provides valuable information regarding protein dynamics within cells and the changes in cell shape and organization. Several techniques have been used to introduce plasmids encoding fluorescently tagged proteins into differentiated podocytes for live imaging, including transfection [Endlich et al.
2009; Welsch et al. 2005], microinjection [Lennon et al. 2008], and electroporation/nucleofection [Soda et al. 2012]. Proteins directly labeled with fluorophores can also be introduced into podocytes by microinjection [Welsch et al. 2005]. Adenoviral infection has also been successfully used to introduce fluorescently labeled constructs encoding actin-binding protein, α-actinin-4, into podocytes [Michaud et al. 2009].
In this paper, we provide a detailed characterization of the use of adenoviral vectors for expression of fluorescently tagged proteins in podocytes
300 Appendix 3 Technique paper for live cell imaging. We test the efficiency of adenoviral infection and show that adenoviral vectors provide a versatile system for reliable protein expression in podocytes without significant toxicity, allowing simultaneous introduction of multiple constructs for multi-color imaging.
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AP 3.4 Reagents and instruments
Microscopes
Epifluorescence imaging was performed using either a Nikon Eclipse TE-
2000E microscope equipped with a PlanApo TIRF 60x/1.45 NA objective,
Harvard Instruments dish warmer for 37°C incubation, and Hamamatsu ORCAII
CCD camera or a Leica AF6000 LX deconvolution microscope equipped with a
40x/1.25 NA and a 63x/1.3 NA HCX PL APO objectives, an environmental chamber, and Andor LucaR EMCCD camera. Confocal imaging was performed using Perkin-Elmer UltraView VoX Spinning Disk Confocal system mounted on a
Nikon Eclipse Ti microscope and equipped with a 60x/1.49 NA APO TIRF objective, Hamamatsu C9100-50 EMCCD camera, and an environmental chamber to maintain cells at 37°C.
Reagents
RPMI 1640 medium 1x (Hyclone), Fetal Bovine Serum (FBS), and antibiotic/antimycotic solution (ABM) (100x) were purchased from Fisher
Sceintific. Rat tail collagen type I and type IV were obtained from BD
Biosciences. Interferon-γ was purchased from EMD Chemicals. Adenovirus kits
(AdEasyTM Adenoviral Vector System and AdEasy Virus Purification Kits) were from Agilent Technologies. Clarity enhanced chemiluminescence substrate for
Western blot analysis was from Bio-Rad.
Mouse anti-synaptopodin antibody (clone G1D4) was from Meridian Life
Science. Rabbit anti-Myo1e antibody was prepared using GST-tagged human
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Myo1e tail as the antigen, and its specificity was characterized using tissue lysates prepared from Myo1e-null and wild type mice.
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AP 3.5 Methods
Podocyte maintenance
Undifferentiated podocytes originally isolated by [Mundel et al. 1997b] were cultured as described therein. Briefly, 1-2 x105 undifferentiated podocytes were plated into a 100 mm collagen I-coated tissue culture dish in 10 ml complete RPMI (RPMI + antibiotic/antimycotic solution + 10% FBS + interferon-γ
o at 50 U/ml) at 33 C, 5% CO2 and subcultured every 2 - 3 days to avoid confluency.
Podocyte differentiation for imaging
1.5 x 104 undifferentiated podocytes were plated into a 35mm collagen IV coated glass bottom dish (Mattek, Ashland, MA) or into a 35mm dish containing collagen-coated glass coverslip in 2 ml complete RPMI without interferon-γ, shifted to 37oC and allowed to differentiate for 14 days. Medium was changed every 2 – 3 days. At the time of infection and imaging, podocytes were at approximately 60-70% confluency.
Collagen coating:
For tissue culture plastic dishes:
Collagen I was diluted to 0.1 mg/ml in PBS with 20 mM acetic acid. 5 ml of diluted collagen I was added to 100mm dish, incubated at 37oC for 1 hour, dishes were washed 3 times with PBS, and used immediately or allowed to dry and stored at 4oC.
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For Mattek glass bottom dishes or coverslips:
Collagen IV was diluted to 10 µg/ml in 0.05 N HCl, added to the glass substrate, incubated at 37oC for 1 hour. After 3 washes with PBS, dishes were used immediately or allowed to dry and stored at 4oC.
Adenoviral vector preparation
AdEasy XL Adenoviral System (Agilent Technologies) was used as the source of the vectors and cells required to clone and package recombinant adenovirus. cDNA encoding protein of interest fused with a fluorescent protein tag (EGFP or mCherry) was initially cloned into the pShuttle vector using either standard restriction/ligation mediated cloning or the InFusion cloning kit
(Clontech). Coding sequences of human Myo1e, human ZO-1 (obtained from
Alan Fanning, UNC), human synaptopodin (long isoform, a generous gift from
Peter Mundel, Mass General) [Asanuma et al. 2005], rat clathrin light chain
(generous gift from Pietro De Camilli, Rushika Perera, and Roberto Zoncu), or
Lifeact were used for cloning into the pShuttle vector. All constructs were expressed under the control of CMV promoter, which was either transferred into the promotorless pShuttle vector from the intermediate cloning vector (for example, pEGFP-C1 (Clontech)) or was contained within the pShuttle-CMV vector. The fusion constructs in pShuttle vector were tested by standard DNA transfection into an appropriate eukaryotic cell line (e.g., Cos-7) to confirm protein expression and localization prior to the investment of the effort required to make adenoviral particles.
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Recombination of the shuttle vector with pAdEasy-1 plasmid, which contains most of the human adenoviral Ad5 genome, was performed according to the manufacturer’s instructions (Agilent Technologies). Viral particles were obtained by transfection of HEK-293 or AD-293 cells. According to the standard protocol, the recombinant pAd DNA needs to be linearized before transfection, yielding a ~30Kb fragment and either a 3 or 4.5 Kb bacterial sequence fragment.
The resulting adenoviral genome fragment (~30 Kb) can be isolated by gel purification. Due to the large size of the DNA fragment, standard resin column gel purification methods may not produce high yield of DNA necessary for productive transfection. We found that phenol:chloroform:isoamyl alcohol extraction of the entire digest mixture followed by ethanol precipitation produced adequate yields of transfectable DNA.
60 mm dish of HEK-293 or AD293 cells was transfected using JetPEI
(Polyplus) and 5 µg of linearized pAd plasmid. Transfected cultures were monitored for cytopathic effect (cells rounding up and floating). When cytopathic effect was evident, cells were scraped off and transferred along with the culture media (~ 5ml) into a 15ml screw top tube. Cells were lysed by three cycles of freezing at -80o C and thawing in a 25oC water bath followed by vigorous vortexing after each cycle. Alternatively, to shorten the time necessary to freeze the lysate, a dry ice / ethanol bath may be used. Cellular debris was removed by centrifugation at 3000 x g at 4oC for 15 minutes. The supernatant, containing the
306 Appendix 3 Technique paper primary crude viral lysate (primary CVL), was transferred into a sterile tube and used to test protein expression and localization in podocytes.
For plaque purification of the virus, 5 x 105 293 cells were plated into each
o well of a 6 well tissue culture plate and incubated at 37 C, 5% CO2 overnight. 0.1 ml of serial dilutions of primary CVL from 10-3 to 10-7 or mock infection control
o were used to infect each well by incubation for 1 hour, 37 C, 5% CO2 with gentle rocking of plate every 15 minutes.
Prior to infection, we prepared 2x DMEM (from powder), 2x ABM, 15%
FBS, warmed up to 37o C. 3% SeaPlaque agarose (Lonza) in water was melted by boiling and cooled to 42o C. Equal volumes of 2x media and agarose solution was mixed and 3 ml of the mixture was immediately but gently overlayed on infected cell monolayers. Cells were maintained at 37oC in 5% CO2. Cultures were overlayed with 1 ml newly prepared media / agarose mixture based on the color change of phenol red indicator to yellow, usually at least once per week.
Plaques became visible within one to three weeks. Several well isolated plaques were collected using a micropipettor or a Pasteur pipette, by aspirating the plaque and dispensing into a microcentrifuge tube containing 300 ul infection media. Cells were lysed by freeze / thawing 3 times, vortexing after each thaw.
Cellular debris and agarose were pelleted by centrifugation at 15,000 x g, 5 minutes, and supernatant (CVL0) was transferred to new tube.
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Amplification 1:
5x105 293 cells were plated into 35mm dish and grown overnight. 100 µl of
CVL0 + 400 µl infection media was added to the monolayer for one hour, with gentle rocking every 15 minutes. After 1 hour, 1.5 ml medium was added to each dish. Cytopathic effect was typically observed after 2 – 3 days. At this point, cells were scraped off, media and cells were collected and transferred to a Falcon
2063 tube. Cells were lysed by freeze / thawing 3 times, vortexing after each thaw. Cellular debris was removed by centrifugation at 3000 x g at 4o C for 15 minutes. The viral supernatant CVL1 was transferred into a sterile Falcon 2063 tube.
Amplification 2:
2 x 106 293 cells were plated into 60mm dish and grown overnight. Using
500 µl CVL1, cell monolayer was infected for one hour, with gentle rocking every
15 minutes. After 1 hour, 2.5 ml medium was added to each dish. Cytopathic effect was typically complete after 2 days. Viral supernatant was collected as described for amplification 1, producing CVL2.
Large Scale Amplification:
The day prior to infection, a 175 cm2 tissue culture flask of 293 cells was split in to five 175 cm2 flasks so that the monolayer was 80% confluent the day of infection. 2.5 ml CVL2 was diluted in 22.5 ml infection media. Each of the 5
o flasks was infected with 5ml diluted CVL2 at 37 C, 5% CO2 for one hour, with gentle rocking every 15 minutes. After 1 hour, 15 ml of medium was added to
308 Appendix 3 Technique paper each flask. When cytopathic effect was complete, cells were scraped and collected for purification of high titer viral stock. AdEasy Virus Purification Kit
(Agilent Technologies) was used to purify high titer viral stocks. Purified virus was exchanged into the cryoprotective long-term storage buffer as suggested by the manufacturer.
Viral concentration (plaque forming units (PFU)/ml) was determined by plaque assay following the same procedure as described above for the isolation of the viral plaques. Generally, 1 ml viral stock of 1 x 108 PFU/ml was obtained from each preparation. Stocks were aliquoted and stored at -80oC
Infecting podocytes
Cells were infected by diluting virus in infection media (RPMI, ABM, 2%
FBS) and adding to cells 24 hrs before imaging. Typical dilution used for live cell imaging was 1 x 105 PFU/ml; however, some viruses were diluted to a lower concentration as empirically determined to avoid overexpression (for example, mCherry-Lifeact). For expression of multiple markers, two viruses were combined and used for infection at the same time. Culture media was removed and replaced with the diluted virus in a minimum volume required to cover the cell monolayer, typically 0.1 ml in a 35 mm dish or 1 ml in a 100 mm dish. Using this infection protocol with viral dilution of 1 x 105 PFU/ml, the multiplicity of infection
(MOI, or the ratio of virus particles to cells) was estimated to be approximately 1.
It should be noted that, based on testing various infection protocols, we did not observe an increase in infection efficiency with increasing volume of virus-
309 Appendix 3 Technique paper containing solution. Thus, infection efficiency appears to depend primarily of virus concentration (PFU/ml) rather than on the total number of virus particles in the dish.
Virus may be added directly to the entire culture volume at the same concentration. However, to minimize the chance of a spill as well as to conserve the viral material, the minimum volume method is preferable. The dishes were
o returned to 37 C, 5% CO2 for 1 hour with gentle rocking every 15 minutes to distribute the diluted virus across the monolayer. At the completion of the infection period, diluted virus was removed if desired and complete media was added to the culture (RPMI, ABM, 10% FBS).
For time-course infection, cells were infected at a concentration of 1 x 106
PFU/ml and transgenes allowed to express for different amounts of time before being lysed and analyzed by western blot.
Western blotting and gel quantitation
WT podocytes were infected with different concentrations of adenoviral vector encoding GFP-Myo1e for 24 hours. The cells were then lysed in ice-cold lysis buffer (50mM Tris-HCl pH=7.5, 150 mM NaCl, 1% Triton X-100, and 10% glycerol) with Complete protease inhibitor (Roche), followed by SDS-PAGE. The samples were blotted using anti-Myo1e antibody, and developed with Clarity enhanced chemiluminescence substrate (Bio-Rad).
For gel quantitation, the images were taken using a gel imager (ChemiDoc system, Bio-Rad) and quantified using ImageJ software.
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Immunofluorescence staining
WT podocytes were plated on collagen IV coated coverslips and processed as described (Chase et al., 2012). The cells were stained for endogenous synaptopodin.
MTT assay for cell viability
Wild type podocytes were plated at 1000 cells/ well in a 96-well plate and allowed to differentiate at 37oC. Cells were mock-infected or infected with the indicated concentrations of the adenoviral vector encoding mCherry (pAd- mCherry) for the indicated time period. To perform the assay, the medium was removed and replaced with 100 µl of DMEM without phenol red and 10 µl of 5 mg/ml MTT (methylthiazolyldiphenyl-tetrazolium bromide), and the plate was incubated 4 hours at 37oC. Following the incubation, 85 µl of the medium were removed and 50 µl of DMSO were added to each well. The plate was incubated at 37oC for 10 min, and the absorbance at 540 nm wavelength was measured using a plate reader.
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AP 3.6 Results
AP 3.6.1 Adenoviral vector infection has high efficiency and low toxicity.
Using adenoviral vectors for infecting differentiated mouse podocytes in culture, we routinely observed high efficiency of infection (more than 50% of cells infected). To quantify the infection efficiency, we used an adenoviral vector encoding mCherry fluorescent protein (Ad-mCherry) to infect both differentiated and non-differentiated podocytes. Four different concentrations of adenovirus
(PFU/ml of medium) were used in the experiments. To collect the images of infected cells, an identical exposure time (1.5 s) was used to capture mCherry fluorescence signal in all cells. More than 90% of cells were infected using adenovirus at 106 PFU/ml, with the efficiency of infection decreasing gradually at lower virus concentration (AP Fig. 3.1A). Slightly higher infection efficiency was observed when applying adenovirus to non-differentiated podocytes compared to the differentiated cells. However, this difference was relatively small, confirming our observations that adenovirus can be used to readily infect differentiated, non- proliferating podocytes. Since mCherry is a relatively small protein (~20kD), we also used adenovirally encoded mCherry-synaptopodin long isoform (~120kD) to test the infection efficiency of a vector containing a larger expression construct.
As shown in AP Fig. 3.1B, the size of the fusion protein did not affect infection efficiency.
Some methods of introducing expression constructs into differentiated cells, such as electroporation, are accompanied by the high degree of cell death.
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Our observations indicated that adenoviral infection was not accompanied by toxic effects on cells, since very little cell detachment and death was observed in the infected cultures. To quantify the effects of adenoviral infection on cell viability, we used an MTT assay (AP Fig. 3.1C-D). We did not observe an increase in cell death in virus-infected cells compared to the mock infection at one day post-infection (AP Fig. 3.1C), and even after 4 days cell number was not decreased in the infected wells compared to the mock-infected cells (AP Fig.
3.1D). Thus, at concentrations used in our experiments, adenoviral infection of podocytes does not affect cell viability.
AP 3.6.2 Expression level of recombinant proteins.
Overexpression of recombinant proteins compared to the endogenous level of the same protein is often a concern in transient transfection experiments, particularly those aimed at analyzing protein dynamics in live cells. In order to compare the expression level of a recombinant protein achieved using either adenoviral infection or a transient DNA tranfection, we used constructs encoding
EGFP-tagged myosin 1e (Ad-GFP-Myo1e or pEGFP-C1-Myo1e). An antibody against Myo1e was used to determine the amount of GFP-Myo1e and the endogenous Myo1e (AP Fig. 3.2). As shown in AP Fig. 3.2A, GFP-Myo1e expression level in podocytes 24 hours after an infection with the adenovirus at
104 or 5 x 104 PFU/ml was 0.8 and 3-fold higher than the level of expression of endogenous Myo1e, respectively. GFP-Myo1e expression level in Cos-7 cells 24 hours after standard DNA transfection was so high that the endogenous Myo1e
313 Appendix 3 Technique paper band was completely obscured by the overexpressed protein (AP Fig. 3.2B).
According to AP Fig. 3.1A, using adenovirus at 104 and 5 x 104 PFU/ml results in infection of approximately 15% and 45% of cells, respectively. When the fraction of cells infected was taken into account, we calculated that GFP-Myo1e concentration in each infected cell was on average 4.6-4.7-fold higher than the level of endogenous Myo1e (AP Fig. 3.2A). While this level of overexpression was more moderate than that observed with the DNA transfection of Cos-7 cells, we decided to further investigate whether expression level of the recombinant protein could be modulated by changing the time of expression. We conducted a time course experiment using the adenoviral vector encoding GFP-Myo1e to infect wild type podocytes at a concentration of 106 PFU/ml, at which more than
95% of cells should be infected. GFP-Myo1e expression level was analyzed at different time points post-infection (AP Fig. 3.2C). At 4 hours post infection, the expression level of GFP fusion protein was similar to the endogenous level of
Myo1e (91% of the endogenous level). Thus, adjusting the time of expression makes it possible to express adenovirally delivered constructs at the same level as the endogenous protein. This observation may be particularly useful in some rescue experiments to avoid potential negative effects of protein overexpression.
AP 3.6.3 Using adenovirally encoded cytoskeletal probes to visualize podocyte cytoskeleton.
We have developed several adenoviral vectors encoding cytoskeletal marker proteins fused to EGFP or mCherry that can be used for live cell imaging.
314 Appendix 3 Technique paper
These proteins (AP Fig. 3.3) represent important markers of the podocyte cytoskeleton and cytoskeleton-associated structures. All of the markers shown can be visualized using either confocal microscopy (spinning disk confocal microscope was used to allow rapid acquisition of images from live cells) or conventional epifluorescence microscopy. Since podocytes are fairly flat, satisfactory results were obtained using both confocal and epifluorescence imaging. All microscopes used in this study were equipped with temperature controlled chambers or stage warmers to maintain cells at 37°C.
Myosin 1e (Myo1e) is an unconventional myosin necessary for normal podocyte development [Chase et al. 2012] and represents one of the protein components of the glomerular slit diaphragm [Bi et al. 2013]. Myo1e is involved in clathrin-dependent endocytosis [Cheng et al. 2012; Krendel et al. 2007] and formation of cell-cell contacts [Bi et al. 2013]. GFP-Myo1e expressed in podocytes localized to the newly forming lamellipodia and filopodia (AP Fig. 3.3A, arrows), to cell-cell junctions, and to clathrin-coated vesicles (AP Fig. 3.5C-D;
[Soda et al. 2012]). Mutations in MYO1E gene are associated with familial kidney disease focal segmental glomerulosclerosis [Al-Hamed et al. 2013; Mele et al.
2011; Sanna-Cherchi et al. 2011]. One of the disease-associated mutations in
MYO1E replaces a highly conserved alanine 159, located near the Switch I region in the motor domain, with proline [Mele et al. 2011]. When adenovirally encoded GFP-Myo1e-A159P was expressed in podocytes, it displayed a diffuse
315 Appendix 3 Technique paper cytoplasmic localization, indicating the loss of the ability to localize to actin- containing structures (AP Fig. 3.3B, [Bi et al. 2013; Mele et al. 2011]).
Lifeact, a 17 amino acid-long peptide derived from a budding yeast actin filament-binding protein ABP140 [Riedl et al. 2008], is a good marker of the dynamic actin cytoskeletal structures since it does not interfere with actin assembly and disassembly [Riedl et al. 2010]. Using adenovirally encoded mCherry-Lifeact, we were able to visualize F-actin in live podocytes (AP Fig.
3.3D, AP Fig. 3.4E-F). Since Lifeact is also a good marker of actin-containing cortical structures, we were able to readily track the forward movement of the leading edge (AP Fig. 3.4G kymograph).
Synaptopodin, an actin-binding protein, is an important regulator of the podocyte actin cytoskeleton [Asanuma et al. 2005; Asanuma et al. 2006; Deller et al. 2003; Faul et al. 2008; Mundel et al. 1997a]. mCherry-synaptopodin decorated actin stress fibers in podocytes, forming a characteristic punctate pattern reminiscent of myofibrils (AP Fig. 3.3C). The localization of mCherry- synaptopodin was similar to that of the endogenous synaptopodin, as revealed by immunostaining (AP Fig. 3.3C’). Because synaptopodin creates a discontinuous, speckle-like pattern of actin labeling, it can be used to trace contraction and lateral movements of stress fibers. AP Fig. 3.4 (A-C) shows confocal images of live podocytes infected with adenovirus encoding mCherry- synaptopodin. Individual frames from a time-lapse movie of mCherry-labeled synaptopodin are shown in AP Fig. 3.4 (B and C). Individual synaptopodin spots
316 Appendix 3 Technique paper
(arrow in 4B) underwent retrograde flow away from the cell edge, which is characteristic of actin-containing structures. Examples of retrograde flow of synaptopodin spots are shown using kymographs (AP Fig. 3.4D). The rate of retrograde flow measured from the kymographs of synaptopodin spots (0.38 +/-
0.24 µm/min, N=4 cells) was similar to the relatively slow rate of retrograde actin flow observed in some fibroblast cell lines [Theriot and Mitchison 1992].
Synaptopodin labeling also allowed visualization of actin stress fiber contraction
(AP Fig. 3.4C). When paired with GFP-tagged Myo1e, synaptopodin can be used to visualize stress fibers while Myo1e highlights the cell edge and is enriched in lamellipodia (AP Fig. 3.5).
Clathrin plays a key role in endocytosis, which is important for podocyte functions [Soda et al. 2012]. Transient expression of fluorescently labeled clathrin light chain using DNA transfection of cell lines such as Cos-7 often results in overexpression and loss of vesicular localization, based on our experience (data not shown). When using an adenoviral vector to introduce clathrin light chain into podocytes, overexpression of clathrin light chain was rarely observed, allowing successful imaging of endocytic vesicles. In AP Fig. 3.3E’, clathrin coated endocytic vesicles can be easily visualized. AP Fig. 3.5 C-D also show the colocalization of clathrin (red) and Myo1e (green) in endocytic vesicles (arrows).
ZO-1 is an actin-associated protein and an important component of the glomerular slit diaphragm [Reiser et al. 2000; Schnabel et al. 1990]. We have previously shown that ZO-1 interacts with Myo1e [Bi et al. 2013]. In AP Fig. 3.3F,
317 Appendix 3 Technique paper mCherry tagged ZO-1 localizes to the cell-cell contacts between podocytes
(arrows).
318 Appendix 3 Technique paper
AP 3.7 Discussion
Imaging of podocyte cytoskeletal organization and dynamics can provide valuable insights into the functioning of the molecular machinery responsible for the complex architecture of the glomerular filtration barrier. Introduction of conditionally immortalized podocyte cell lines represented a key advancement in the ability to study podocyte cell biology in vitro [Shankland et al. 2007].
Following the development of podocyte cell lines, live imaging of podocyte cytoskeletal structures has been used to characterize such dynamic processes as formation of actin-containing protrusions [Akilesh et al. 2011], actin cytoskeletal reorganization in response to the treatment with growth factors and serum components [Endlich et al. 2009; Lennon et al. 2008], contraction of stress fibers and reorganization of focal adhesions [Endlich et al. 2009], and movement and internalization of endocytic vesicles and endosomal compartments associated with actin and other cytoskeletal proteins [Soda et al. 2012; Welsch et al. 2005]. Fluorescently labeled cytoskeletal proteins have also been utilized for measurements of protein turnover in podocytes by fluorescence recovery after photobleaching (FRAP) [Endlich et al. 2009].
Since transient transfection of podocytes is characterized by relatively low efficiency, the use of viral transduction for expression of fluorescently tagged proteins represents a promising technique for the studies of podocyte cell biology. Both lentiviral and adenoviral vectors have been used for podocyte transduction. The lentiviral and retroviral vectors are able to integrate into the
319 Appendix 3 Technique paper host cell genome, resulting in stable transfection. This property is beneficial for the production of stable cell lines but could also result in insertional mutagenesis by disrupting or activating host cell genes. On the other hand, adenoviral vectors offer a highly efficient system for short-term protein expression, without the likelihood of disrupting endogenous gene expression.
Adenoviruses have been reported to cause changes in the cytoskeletal organization of host cells upon infection, due to activation of a number of cell signaling pathways [Taylor et al. 2011]. These effects of adenoviruses have been mapped primarily to the genes located in the E1 region. The E1 region has been removed from the adenoviruses engineered for use in laboratory studies, such as those produced using the AdEasy system. Removal of the E1 region renders adenovirus replication-incompetent so that infectious adenoviral particles can be produced only in special packaging cells, such as HEK-293, which supply the E1 gene products. This modification not only increases the safety of adenoviral vectors but also reduces the chances of adenoviral infection causing reorganization of the host cell cytoskeleton. However, in every experiment using adenoviral vectors, a possibility of the viral infection inducing changes in the actin organization should be considered, and careful use of controls (such as infecting cells with an adenovirus encoding GFP or mCherry followed by examination of actin organization in infected and non-infected cells) may be advisable.
Recent advancements in the field of podocyte biology include development of the techniques that allow fluorescent labeling of glomerular
320 Appendix 3 Technique paper podocytes in vivo using transgenic mice [Grgic et al. 2012; Hohne et al. 2013] and zebrafish [He et al. 2011] that express genetically encoded fluorescent labels in a podocyte-specific manner. In future studies, these cell type-specific fluorescent labels can be combined with the intravital imaging of renal glomeruli by multi-photon microscopy [Peti-Peterdi and Sipos 2010] to allow the direct observation of dynamic rearrangements of podocyte foot processes and changes in glomerular permeability in vivo.
Development of the techniques that allow efficient labeling of podocytes with specific fluorescent markers would further broaden the toolkit of the researchers interested in the cell biology of podocytes and allow the dissection of molecular pathways leading to the development of glomerular disorders. As described in this paper, adenoviral vectors encoding fluorescently tagged cytoskeletal proteins provide another valuable tool that can be used to analyze podocyte cytoskeletal dynamics. Our study shows that adenoviral vectors can be used to introduce fluorescently tagged cytoskeletal markers into differentiated cultured podocytes with high efficiency and low toxicity. Viral concentration and infection time can be adjusted to regulate the level of overexpression of the protein of interest, and multiple fluorescent markers can be combined in the same cell. In the future, this technique may be adapted to allow fluorescent labeling of podocytes in vivo for intravital imaging of glomerular dynamics.
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AP 3.8 Reference
Akilesh S, Suleiman H, Yu H, Stander MC, Lavin P, Gbadegesin R, Antignac C, Pollak M, Kopp JB, Winn MP and others. 2011. Arhgap24 inactivates Rac1 in mouse podocytes, and a mutant form is associated with familial focal segmental glomerulosclerosis. J Clin Invest 121(10):4127-37. Al-Hamed MH, Al-Sabban E, Al-Mojalli H, Al-Harbi N, Faqeih E, Al Shaya H, Alhasan K, Al-Hissi S, Rajab M, Edwards N and et al.. 2013. A molecular genetic analysis of childhood nephrotic syndrome in a cohort of Saudi Arabian families. J Hum Genet. Asanuma K, Kim K, Oh J, Giardino L, Chabanis S, Faul C, Reiser J, Mundel P. 2005. Synaptopodin regulates the actin-bundling activity of alpha-actinin in an isoform-specific manner. J Clin Invest 115(5):1188-98. Asanuma K, Yanagida-Asanuma E, Faul C, Tomino Y, Kim K, Mundel P. 2006. Synaptopodin orchestrates actin organization and cell motility via regulation of RhoA signalling. Nat Cell Biol 8(5):485-91. Bi J, Chase SE, Pellenz CD, Kurihara H, Fanning AS, Krendel M. 2013. Myosin 1e is a component of the glomerular slit diaphragm complex that regulates actin reorganization during cell-cell contact formation in podocytes. Am J Physiol Renal Physiol 305(4):F532-44. Brown EJ, Schlondorff JS, Becker DJ, Tsukaguchi H, Uscinski AL, Higgs HN, Henderson JM, Pollak MR. 2010. Mutations in the formin gene INF2 cause focal segmental glomerulosclerosis. Nat Genet 42(1):72-6. Chase SE, Encina CV, Stolzenburg LR, Tatum AH, Holzman LB, Krendel M. 2012. Podocyte-specific knockout of myosin 1e disrupts glomerular filtration. Am J Physiol Renal Physiol 303(7):F1099-106. Cheng J, Grassart A, Drubin DG. 2012. Myosin 1E coordinates actin assembly and cargo trafficking during clathrin-mediated endocytosis. Mol Biol Cell 23(15):2891-904. Deller T, Korte M, Chabanis S, Drakew A, Schwegler H, Stefani GG, Zuniga A, Schwarz K, Bonhoeffer T, Zeller R and others. 2003. Synaptopodin-deficient mice lack a spine apparatus and show deficits in synaptic plasticity. Proc Natl Acad Sci U S A 100(18):10494-9. Endlich N, Schordan E, Cohen CD, Kretzler M, Lewko B, Welsch T, Kriz W, Otey CA, Endlich K. 2009. Palladin is a dynamic actin-associated protein in podocytes. Kidney Int 75(2):214-26. Faul C, Asanuma K, Yanagida-Asanuma E, Kim K, Mundel P. 2007. Actin up: regulation of podocyte structure and function by components of the actin cytoskeleton. Trends Cell Biol 17(9):428-37. Faul C, Donnelly M, Merscher-Gomez S, Chang YH, Franz S, Delfgaauw J, Chang JM, Choi HY, Campbell KN, Kim K and others. 2008. The actin cytoskeleton of kidney podocytes is a direct target of the antiproteinuric effect of cyclosporine A. Nat Med 14(9):931-8. Greka A, Mundel P. 2012. Cell biology and pathology of podocytes. Annu Rev Physiol 74:299-323. Grgic I, Brooks CR, Hofmeister AF, Bijol V, Bonventre JV, Humphreys BD. 2012. Imaging of podocyte foot processes by fluorescence microscopy. J Am Soc Nephrol 23(5):785- 91. He B, Ebarasi L, Hultenby K, Tryggvason K, Betsholtz C. 2011. Podocin-green fluorescence protein allows visualization and functional analysis of podocytes. J Am Soc Nephrol 22(6):1019-23. Hohne M, Ising C, Hagmann H, Volker LA, Brahler S, Schermer B, Brinkkoetter PT, Benzing T. 2013. Light microscopic visualization of podocyte ultrastructure demonstrates oscillating glomerular contractions. Am J Pathol 182(2):332-8. Jat PS, Noble MD, Ataliotis P, Tanaka Y, Yannoutsos N, Larsen L, Kioussis D. 1991. Direct derivation of conditionally immortal cell lines from an H-2Kb-tsA58 transgenic mouse. Proc Natl Acad Sci U S A 88(12):5096-100.
322 Appendix 3 Technique paper
Kaplan JM, Kim SH, North KN, Rennke H, Correia LA, Tong HQ, Mathis BJ, Rodriguez-Perez JC, Allen PG, Beggs AH and others. 2000. Mutations in ACTN4, encoding alpha-actinin-4, cause familial focal segmental glomerulosclerosis. Nat Genet 24(3):251-6. Katsuya K, Yaoita E, Yoshida Y, Yamamoto Y, Yamamoto T. 2006. An improved method for primary culture of rat podocytes. Kidney Int 69(11):2101-6. Kaufman L, Yang G, Hayashi K, Ashby JR, Huang L, Ross MJ, Klotman ME, Klotman PE. 2007. The homophilic adhesion molecule sidekick-1 contributes to augmented podocyte aggregation in HIV-associated nephropathy. FASEB J 21(7):1367-75. Kim JM, Wu H, Green G, Winkler CA, Kopp JB, Miner JH, Unanue ER, Shaw AS. 2003. CD2-associated protein haploinsufficiency is linked to glomerular disease susceptibility. Science 300(5623):1298-300. Krendel M, Osterweil EK, Mooseker MS. 2007. Myosin 1E interacts with synaptojanin- 1 and dynamin and is involved in endocytosis. FEBS Lett 581(4):644-50. Lennon R, Singh A, Welsh GI, Coward RJ, Satchell S, Ni L, Mathieson PW, Bakker WW, Saleem MA. 2008. Hemopexin induces nephrin-dependent reorganization of the actin cytoskeleton in podocytes. J Am Soc Nephrol 19(11):2140-9. Mathieson PW. 2012. The podocyte as a target for therapies--new and old. Nat Rev Nephrol 8(1):52-6. Mele C, Iatropoulos P, Donadelli R, Calabria A, Maranta R, Cassis P, Buelli S, Tomasoni S, Piras R, Krendel M and others. 2011. MYO1E mutations and childhood familial focal segmental glomerulosclerosis. N Engl J Med 365(4):295-306. Michaud JL, Hosseini-Abardeh M, Farah K, Kennedy CR. 2009. Modulating alpha- actinin-4 dynamics in podocytes. Cell Motil Cytoskeleton 66(3):166-78. Mundel P, Heid HW, Mundel TM, Kruger M, Reiser J, Kriz W. 1997a. Synaptopodin: an actin-associated protein in telencephalic dendrites and renal podocytes. J Cell Biol 139(1):193-204. Mundel P, Reiser J. 2010. Proteinuria: an enzymatic disease of the podocyte? Kidney Int 77(7):571-80. Mundel P, Reiser J, Zuniga Mejia Borja A, Pavenstadt H, Davidson GR, Kriz W, Zeller R. 1997b. Rearrangements of the cytoskeleton and cell contacts induce process formation during differentiation of conditionally immortalized mouse podocyte cell lines. Exp Cell Res 236(1):248-58. Ni L, Saleem M, Mathieson PW. 2012. Podocyte culture: tricks of the trade. Nephrology (Carlton) 17(6):525-31. Pavenstadt H, Kriz W, Kretzler M. 2003. Cell biology of the glomerular podocyte. Physiol Rev 83(1):253-307. Peti-Peterdi J, Sipos A. 2010. A high-powered view of the filtration barrier. J Am Soc Nephrol 21(11):1835-41. Reiser J, Kriz W, Kretzler M, Mundel P. 2000. The glomerular slit diaphragm is a modified adherens junction. J Am Soc Nephrol 11(1):1-8. Reiser J, Sever S. 2013. Podocyte biology and pathogenesis of kidney disease. Annu Rev Med 64:357-66. Reiser J, von Gersdorff G, Loos M, Oh J, Asanuma K, Giardino L, Rastaldi MP, Calvaresi N, Watanabe H, Schwarz K and others. 2004. Induction of B7-1 in podocytes is associated with nephrotic syndrome. J Clin Invest 113(10):1390-7. Riedl J, Crevenna AH, Kessenbrock K, Yu JH, Neukirchen D, Bista M, Bradke F, Jenne D, Holak TA, Werb Z and others. 2008. Lifeact: a versatile marker to visualize F-actin. Nat Methods 5(7):605-7. Riedl J, Flynn KC, Raducanu A, Gartner F, Beck G, Bosl M, Bradke F, Massberg S, Aszodi A, Sixt M and others. 2010. Lifeact mice for studying F-actin dynamics. Nat Methods 7(3):168-9.
323 Appendix 3 Technique paper
Saleem MA, O'Hare MJ, Reiser J, Coward RJ, Inward CD, Farren T, Xing CY, Ni L, Mathieson PW, Mundel P. 2002. A conditionally immortalized human podocyte cell line demonstrating nephrin and podocin expression. J Am Soc Nephrol 13(3):630-8. Sanna-Cherchi S, Burgess KE, Nees SN, Caridi G, Weng PL, Dagnino M, Bodria M, Carrea A, Allegretta MA, Kim HR and others. 2011. Exome sequencing identified MYO1E and NEIL1 as candidate genes for human autosomal recessive steroid-resistant nephrotic syndrome. Kidney Int 80(4):389-96. Schiwek D, Endlich N, Holzman L, Holthofer H, Kriz W, Endlich K. 2004. Stable expression of nephrin and localization to cell-cell contacts in novel murine podocyte cell lines. Kidney Int 66(1):91-101. Schnabel E, Anderson JM, Farquhar MG. 1990. The tight junction protein ZO-1 is concentrated along slit diaphragms of the glomerular epithelium. J Cell Biol 111(3):1255-63. Shankland SJ, Pippin JW, Reiser J, Mundel P. 2007. Podocytes in culture: past, present, and future. Kidney Int 72(1):26-36. Soda K, Balkin DM, Ferguson SM, Paradise S, Milosevic I, Giovedi S, Volpicelli- Daley L, Tian X, Wu Y, Ma H and others. 2012. Role of dynamin, synaptojanin, and endophilin in podocyte foot processes. J Clin Invest 122(12):4401-11. Taylor MP, Koyuncu OO, Enquist LW. 2011. Subversion of the actin cytoskeleton during viral infection. Nat Rev Microbiol 9(6):427-39. Theriot JA, Mitchison TJ. 1992. Comparison of actin and cell surface dynamics in motile fibroblasts. J Cell Biol 119(2):367-77. Welsch T, Endlich N, Gokce G, Doroshenko E, Simpson JC, Kriz W, Shaw AS, Endlich K. 2005. Association of CD2AP with dynamic actin on vesicles in podocytes. Am J Physiol Renal Physiol 289(5):F1134-43. Welsh GI, Saleem MA. 2012. The podocyte cytoskeleton--key to a functioning glomerulus in health and disease. Nat Rev Nephrol 8(1):14-21.
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AP Figure 3.1. Adenoviral infection of podocytes is characterized by high efficiency and low toxicity.
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AP Figure 3.1. Adenoviral infection of podocytes is characterized by high efficiency and low toxicity. A, adenoviral infection efficiency of non- differentiated vs. differentiated podocytes was determined using different concentrations of mCherry-encoding adenovirus. Fraction of infected cells was determined using a ratio of the number of cells expressing mCherry to the number of DAPI-labeled nuclei. B, adenoviral infection efficiency of differentiated podocytes infected with an adenoviral vector encoding mCherry vs. an adenoviral vector encoding mCherry-synaptopodin. Fraction of infected cells was determined as in A. C, MTT-based viability assay on differentiated podocytes 24 hours after being infected with an adenoviral vector encoding mCherry. Equal numbers of cells were plated into each well and infected with the adenovirus at indicated concentrations. OD540 is proportionate to the number of viable cells. D.
MTT-based viability assay on differentiated podocytes at 96 hours (4 days) after being infected with adenovirus at indicated concentrations.
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AP Figure 3.2. Western blot analysis of cell lysates prepared from differentiated podocytes infected with different concentrations of the adenoviral vector encoding GFP-Myo1e.
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AP Figure 3.2. Western blot analysis of cell lysates prepared from differentiated podocytes infected with different concentrations of the adenoviral vector encoding GFP-Myo1e. A, wild type podocytes were infected with adenoviral vector encoding GFP-Myo1e at indicated concentrations. Cells were lysed at 24 hours post-infection, and gel samples were processed for SDS-
PAGE and Western blot analysis using anti-Myo1e antibody. Ratio of GFP- tagged myosin to the endogenous myosin was calculated by dividing GFP fusion band intensity expression (upper band) by the intensity of the endogenous
Myo1e (lower band). Fold overexpression was calculated by taking into account the fraction of cells that are expected to be infected at a given adenovirus concentration. B, Cos-7 cells were transfected with GFP-Myo1e plasmid. Cells were lysed 24 hours after the transfection and analyzed by Western blotting. C, wild type podocytes were infected with the adenoviral vector encoding GFP-
Myo1e at the concentration of 106 PFU/ml, which results in infection of >95% of cells. Cells were lysed every 4 hours post infection, and gel samples were analyzed by Western blotting.
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AP Figure 3.3. Localization of fluorescently labeled cytoskeletal markers in differentiated podocytes.
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AP Figure 3.3. Localization of fluorescently labeled cytoskeletal markers in differentiated podocytes. A, B, single confocal images showing localization of
GFP-tagged wild type Myo1e (A) and disease-associated Myo1e mutant (B). A, arrows indicate Myo1e localization to filopodia-like projections and vesicle-bound
Myo1e. B, Myo1e mutant (A159P) does not localize to actin-containing projections or to vesicles. C-F, epifluorescence images showing localization of mCherry-tagged synaptopodin (C), Lifeact (D), Clathrin Light Chain (E), and ZO-1
(F) in differentiated podocytes. C’, immunostaining against endogenous synaptopodin in a differentiated podocyte. E’: enlarged image of the boxed region in E. Arrows in F point to cell-cell junctions, where ZO-1 is enriched. Viral concentrations used to infect podocytes were 105 PFU/ml for all constructs. Scale bars: 10 µm.
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AP Figure 3.4. Visualization of podocyte cytoskeletal dynamics using live- cell imaging.
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AP Figure 3.4. Visualization of podocyte cytoskeletal dynamics using live- cell imaging. Differentiated mouse podocytes were infected with either mCherry- tagged synaptopodin (A-D, confocal images) or mCherry-tagged Lifeact (E-G, epifluorescence images). B and C, enlarged images of the boxed regions in A at the indicated time points. Arrow in B points to a synaptopodin-labeled structure undergoing retrograde flow away from the cell edge. Arrows in C, pointing to synaptopodin speckles within stress fibers, illustrate stress fiber contraction
(decreasing distance between the arrows over time). D, kymographs generated using red lines in B illustrate retrograde flow of synaptopodin-labeled actin bundles. F, a time-lapse sequence showing enlarged images of the boxed region in E. G, kymograph generated based on the time-lapse sequence shown in F illustrates protrusion of the cell edge. Scale bar: 10 µm.
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AP Figure 3.5. Visualization of podocyte cytoskeletal dynamics using multi-color live cell imaging.
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AP Figure 3.5. Visualization of podocyte cytoskeletal dynamics using multi- color live cell imaging. Differentiated podocytes were co-infected with GFP-
Myo1e (A) and mCherry-synaptopodin (A’) (A-B), or GFP-Myo1e (C) and mCherry-clathrin light chain (C’) (C-D) and imaged using epifluorescence microscopy. Arrows in A point to synaptopodin-labeled stress fibers, arrows in B indicate Myo1e localization to lamellipodia, arrows in D indicate colocalization of
Myo1e and clathrin on endocytic vesicles. Scale bar: 10 µm.
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