STRUCTURAL AND BIOCHEMICAL ANALYSIS OF THE DYNEIN MOTOR
By: Stephanie A Ketcham
A dissertation submitted to Johns Hopkins University in conformity with the requirements for the degree of Doctor of Philosophy
Baltimore, Maryland October 2016
Copyright © 2016 Stephanie A Ketcham All Rights Reserved
Abstract:
The cytoplasmic dynein motor, hereafter dynein, is necessary for a wide range of subcellular activities. Functions range from the movement of vesicles
(referred to as “low-load” transport) to organization of mitotic spindle poles, where several motors work cooperatively as an ensemble (“high-load” transport).
These different modes are thought to depend on extrinsic regulatory components.
The first such factor to be discovered, dynactin, was identified biochemically as an activator of dynein motility. Later, dynactin was genetically linked with the dynein pathway through mutational screens in several organisms. Additionally, these screens found the dynein regulators NudE and Lis1 (NudF), which contribute to the formation of a high-force, stalled motor.
Despite what is known about the functional roles these factors play in different types of dynein-based motility, high-resolution structural information of dynein complexed with its regulators has been limited. My work lends new insight into the formation of the different dynein motor complexes. In Chapter
Two, I explore the high-force generating motor by analyzing the binding of the
NudE homolog, NudEL, with dynein, and how this interaction enhances Lis1 recruitment. In Chapter Three, I discuss novel insights into the structures of dynein, dynactin and the dynein-dynactin supercomplex bound to microtubules, that were obtained in collaboration with Gabriel Lander and Saikat Chowdhury, in the Department of Integrative and Computational Biology at the Scripps
Research Institute, La Jolla, CA. In Chapter Four, I expand our knowledge of the dynactin structure by biochemically mapping protein-protein interactions within
ii the shoulder subdomain of dynactin, where high-resolution structural information is not available. The work described here contributes to several testable models regarding regulation and loading mechanisms for the dynein motor.
Advisor/Reader:
Trina A Schroer
Professor, Department of Biology, Krieger School of Arts and Sciences, The
Johns Hopkins University
Reader:
M. Andrew Hoyt
Professor, Department of Biology, Krieger School of Arts and Sciences, The
Johns Hopkins University
iii Acknowledgements:
I would like to thank the following people for their help and support throughout the time that this dissertation work was performed.
First, I am greatly appreciative of all the support and guidance I have received from Dr. Trina Schroer. The work presented here was often challenging and/or confusing, and was the cause of much frustration. Despite this, Trina always had faith in my abilities and helped to keep me moving forward. Without her support I would not have completed this work.
I would also like to thank my thesis committee: Dr. Doug Barrick, Dr. Vince
Hilser and Dr. Yixian Zheng. Their suggestions regarding our analysis and methodologies were always appreciated and taken into account. Also, Dr. Andy
Hoyt, for stepping in as a thesis reader when my committee was unable to do so.
I am also grateful to my many collaborators, including Dr. Gabriel Lander and Dr. Saikat Chowdhury (The Scripps Research Institute), without whom we would have been unable to obtain the high-resolution EM structures that facilitated our understanding of the dynein motor. Also, Dr. Yixian Zheng and Dr.
Shusheng Wang (The Carnegie Institute) for their work with us to understand one modality of dynein function.
I must also thank the members of the Schroer Lab. First, Dr. Ting-Yu Yeh, who has been a member of the lab throughout the entirety of my training, his enthusiasm and support has been endless. Also, Dr. Seamus Levine-Wilkinson and Dr. Frances Cheong for their help early on during this work. Their knowledge and advice was greatly appreciated. Finally, Dr. Brett Scipioni and
iv Kevin Delong who supported me during both the ups and downs of graduate school and made the lab a fun place to be.
I am also grateful to all of my friends at JHU and in Baltimore for knowing when to distract me from my work or to allow me to push on. The many conversations, new hobbies and fun times helped to make this experience more enjoyable and will not be forgotten.
Finally I must thank my family. My parents for their continued support through everything I have done, the opportunities they provided and for never letting me feel as if there was anything I could not accomplish. As well as my sisters, Kelli and Caitlin, for both their support and the competitive nature we bring to every aspect of our lives. It was this competitive nature and the persistence that comes with it that helped me to push through many challenges.
v Table of Contents:
Abstract ...... i
Acknowledgements ...... iii
Chapter One - General Introduction...... 1
Subcellular dynein functions ...... 2
Cytoplasmic dynein structure, composition and the dynein cross-bridge
cycle ...... 4
Identification of the “dynein genetic pathway” ...... 8
Conformational regulation of dynein activity ...... 13
Extrinsic regulation of cytoplasmic dynein ...... 16
The stalled, high-force generating motor complex: dynein, NudE and
Lis1 ...... 18
Discovery and structure of the dynein-activating complex, dynactin ...... 23
Stabilization of the ultra-processive dynein-dynactin-scaffold complex ...... 27
Chapter Two - Examining the loading mechanism of the stalled
dynein-NudE(L)-Lis1 motor complex ...... 30
Introduction ...... 31
Results ...... 39
The dynein holo-complex differentially interacts with NudE(L)
fragments...... 39
Dynein IC interacts with NudE(L) fragments, but not full-length
vi NudE(L)...... 42
Lis1 require NudEL to bind the dynein holo-complex ...... 46
Discussion...... 50
Materials and Methods...... 52
Sf9 expression and purification of Lis1 ...... 52
Expression and purification of NudEL, p150Glued and DIC fragments ...... 53
Dynein purification from bovine brain ...... 54
Dynein cosedimentation assay ...... 55
Size exclusion chromatography analysis of protein-protein
interactions...... 56
Chapter Three - EM analysis of the dynein dynactin and dynein-
dynactin-Bicaudal D2 complexes ...... 57
Introduction ...... 58
Results ...... 61
Preparation of chick embryo and bovine brain dynein and dynactin ...... 61
EM analysis of the dynactin complex...... 68
EM analysis of the dynein complex ...... 77
Generation of Dynein-Dynactin-Bicaudal D2 complexes...... 86
Discussion...... 89
Comparative analysis of concurrently generated EM structures ...... 89
Dynactin comparison...... 93
Dynein comparison...... 97
Dynein-dynactin-Bicaudal D2 complex analysis...... 100
vii Structural implications into the regulatory mechanisms of the dynein
motor...... 100
Materials and Methods...... 106
Chick embryo brain dynein and dynactin purifications...... 106
Bovine brain dynein and dynactin purification ...... 108
Bicaudal D2 fragment purification...... 109
DDB complex assembly...... 110
Chapter Four - Mapping the dynactin shoulder domain aimed at
understanding its three-helix bundle structure ...... 112
Introduction ...... 113
Results ...... 115
Dynamitin and p24 sequence analysis ...... 115
Solubilization of p24 fragments with full length dynamitin ...... 121
Dynamitin’s N-terminus (AA 1-212) is not necessary for complex
formation...... 121
Dynamitin’s C-terminus is necessary for complex formation ...... 127
Discussion...... 130
Materials and Methods...... 135
Cloning...... 135
Protein purification ...... 135
Refolding Complexes and complex purification...... 137
Size exclusion chromatography analysis...... 138
Chapter Five - General Conclusions...... 141
viii Predicting the structure of the dynein-dynactin-scaffold complexes ...... 147
Transforming the dynein-Lis1 motor into a motile dynein-dynactin motor
complex...... 148
Concluding remarks ...... 153
Appendix I - Mycalolide B disrupts the dynactin complex...... 154
Introduction ...... 155
Results ...... 159
Mycalolide B depolymerizes actin and destabilizes dynactin ...... 159
Mycalolide B disrupts the dynactin complex in vitro and in vivo ...... 163
Discussion...... 163
Materials and Methods...... 167
Purification of dynein and dynactin ...... 167
HEK293T cell culture ...... 168
Actin polymerization...... 168
Sucrose density gradient analysis ...... 169
Appendix II - Analysis of recombinant p150Glued fragments...... 171
Introduction ...... 172
Results ...... 176
p150Glued fragment design...... 176
Solubility analysis of p150Glued fragments ...... 177
Discussion...... 183
Materials and Methods...... 185
Cloning...... 185
ix Protein purification ...... 186
Refolding complexes ...... 187
Size exclusion chromatography analysis...... 187
Appendix III - The p27 free pool is Rab11a sensitive ...... 191
Introduction ...... 192
Results ...... 196
Transferrin receptor trafficking in cells overexpressing Rab11a
mutants ...... 196
Rab11a overexpression alters the localization of the p27 cluster...... 199
Discussion...... 202
Materials and Methods ...... 202
Rab11a constructs...... 202
Transfections ...... 202
Transferrin uptake assay ...... 203
27A Immunofluorescence ...... 204
Immunofluorescence Microscopy ...... 204
References ...... 205
Curriculum vitae ...... 218
x
Figures and tables:
Figure 1-1. Dynein composition, subunit organization and “step”...... 5
Figure 1-2. Phenotypes of dynein pathway mutants...... 9
Figure 1-3. Dynein and kinesin utilize different autoinhibitory mechanisms...... 14
Figure 1-4. Spindle focusing defects observed in the X. laevis egg extract
system...... 19
Figure 1-5 Loading of the dynein-NudE(L)-Lis1 complex...... 21
Figure 1-6. Dynactin composition and subunit organization...... 24
Figure 2-1. The dynein motor is required for proper spindle formation in
vitro...... 34
Figure 2-2. Recombinant protein fragments...... 37
Figure 2-3. Dynein forms a stable complex with the coiled-coil domain of
NudEL...... 40
Figure 2-4. DIC interacts with the NudEL fragments but not full-length
NudEL...... 43
Figure 2-5. Dynein-Lis1-NudEL form a ternary complex...... 47
Figure 3-1. Negative stained averaged dynein and dynactin particles...... 63
Figure 3-2. Modified bovine brain dynein/dynactin purification protocol...... 66
Figure 3-3. Analysis of dynein exposed to elevated temperatures...... 69
Figure 3-4. Cryo-EM materials...... 72
Figure 3-5. Cryo-EM Arp filament structure...... 75
Figure 3-6. Reconstruction of the dynactin complex structure...... 78
xi Figure 3-7. Focused classification processing procedure...... 81
Figure 3-8. Dynein structure...... 84
Figure 3-9. Generating dynein-dynactin-BicD2 complexes in solution...... 88
Figure 3-10. Generating microtubule bound DDB complexes...... 91
Figure 3-11. Cryo-EM dynactin structure...... 94
Figure 3-12. Structure of the dynein tail...... 98
Figure 3-13. Structure of the dynein tail-dynactin-BicD2 complex...... 101
Figure 3-14. Loading model of the dynein-dynactin-scaffold protein
supercomplex...... 104
Figure 4-1. Sequence analysis of dynamitin and p24...... 117
Figure 4-2. Dynamitin and p24 fragments...... 119
Figure 4-3. The p24 N-terminal 32 amino acids inhibit complex formation
with dynamitin...... 122
Figure 4-4. The N-terminal 212 amino acids expendable for formation of
dynamitin-p24 complexes...... 125
Figure 4-5. The C-terminal 97 amino acids are necessary for dynamitin-p24
complex formation...... 128
Figure 4-6 Structure and polypeptide identity of the shoulder domain ...... 131
Table 4-1. PCR primers used to generate p24 and dynamitin fragments...... 139
Figure 5-1. EM structures of dynein and dynactin ...... 144
Figure 5-2. Transformation of the “stalled” dynein-NudE(L)-Lis1 complex into the
processive dynein-dynactin motor...... 149
Figure A1-1. Mycalolide B binding site on actin and Arp1...... 156
xii Figure A1-2. Mycalolide B depolymerizes actin and disrupts the dynactin
complex...... 160
Figure A1-3. Mycalolide B disruption of purified dynactin and dynactin in
HEK293T cell lysates indicate that the presence of actin acts as an
inhibitor...... 164
Figure A2-1. Dynactin structure...... 174
Figure A2-2. Structural prediction of p150Glued domains used to design
recombinant fragments...... 178
Table A2-1. Analysis of p150Glued fragments ...... 180
Table A2-2. PCR primers used to generate p150Glued fragments...... 189
Figure A3-1. p27 localization is cell cycle dependent...... 193
Figure A3-2. A dominant negative Rab11a mutant alters transferrin
trafficking...... 197
Figure A3-3. p27 localization is altered in Cos7 cells overexpressing ...... Rab11a
mutants...... 200
xiii
Chapter One
General Introduction
1 Subcellular dynein functions:
Proper function and survival of eukaryotic cells requires coordination of many cellular processes, including transport of materials throughout the cell and segregation of intracellular structures during cell division. To perform these broad and essential tasks and maintain organization of intracellular compartments, cells utilize specialized classes of proteins that compose the cytoskeleton. These classes include microtubules, actin and several types of intermediate filaments, all of which play roles in cellular structure. Microtubules and actin also provide polarized tracks for transport 10-12. The subunits of microtubules, α/β tubulin dimers, and globular actin have distinct ends with differing properties, yielding polarized filaments. The plus-ends of both types of filaments are highly dynamic, undergoing rapid growth and shrinkage, whereas the minus-ends are more stable 13,14. The structural polarity of filaments provides directional cues for motor proteins, which are ATPase mechano-enzymes that provide the driving force behind material transport 15-17
Microtubule-based motors fall into two distinct classes. Kinesin family members are primarily plus-end directed motors 18-20, whereas minus-end directed motility relies on dyneins 21-23. The dynein family consists of two classes, axonemal dyneins and cytoplasmic dynein. Axonemal dyneins generate force by sliding microtubule doublets relative to one another within the axoneme to power whole cell motility via ciliary bending 21,24. By contrast, cytoplasmic dynein and motors of the kinesin family are responsible for subcellular transport
2 of a wide array of cargoes, including RNA, proteins, and nearly all membrane bound organelles.
Localization of RNAs and proteins is crucial during embryonic development, where gradients of specific RNAs or proteins result in axial determination in the organism 25-27. At the cellular level, polarity and the plane of cell division is determined by the positioning of RNAs and proteins relative to the microtubule network 5,25,28-30.
Dynein and kinesins play additional key roles during cell division. At the
G2/M transition, the nuclear envelope is broken down. This process is facilitated by dynein, which accumulates on the nuclear membrane and exerts tension on the membrane by moving towards the minus-ends of microtubules on separating nascent spindle poles 31-33. Dynein and kinesins also function within the mitotic spindle, generating forces that allow chromosome capture, alignment and separation prior to cytokinesis 34-36. The final steps of cell division require membrane dynamics at the site of cytokinesis, which is achieved via dynein- based vesicle transport. 37-40. The diversity of functions dynein provides during a single short part of the cell cycle underscores the importance of understanding how the motor functions mechanistically and switches between high and low-load transport.
3 Cytoplasmic dynein structure, composition and the dynein cross-bridge cycle:
The core of the cytoplasmic dynein motor is dynein itself, a homodimeric assembly with a total mass of 1.5 MDa. It comprises two copies each of a heavy chain (HC), intermediate chain (IC), light intermediate chain (LIC) and three distinct light chains (LC7, LC8 and TcTex; Figure 1-1 A & B) 41-46. Each 520 kDa heavy chain contains distinct functional domains, including six AAA+ ATPase-like domains, a microtubule binding stalk, and a tail domain, which is the binding site for the other dynein subunits (Figure 1-1 B & D) 47-49.
Unlike canonical AAA+ ATPases, which are oligomeric assemblies, generally homo-hexamers, the dynein heavy chain includes its six AAA+ domains in a single polypeptide, HC AA 1925-4319. Another unique feature is that
ATPase binding and hydrolytic activity of the six AAA+ motifs differs. AAA1-4 can bind adenosine nucleotide, with AAA1 and 3 able to hydrolyze ATP to ADP, whereas AAA5-6 cannot bind or hydrolyze nucleotide (Figure 1-1 C) 47. ATP hydrolysis at AAA1 appears to be largely responsible for structural changes in the motor domain that underlie changes in microtubule affinity and force generation 50-53. The localized conformational change that accompanies ATP hydrolysis and product release at AAA1 is transmitted through the AAA+ ring to yield a shift in registry of an antiparallel coiled-coil (the “stalk”) that projects from the ring between AAA domains 4 and 5 (Figure 1-1 B & D). This results in a change in the structure of the microtubule-binding domain that alters its affinity for microtubules 54,55.
4
5 Figure 1-1. Dynein composition, subunit organization and “step”. (A) Coomassie
Blue stained SDS-Polyacrylamide gel of purified bovine dynein 3. Subunit labels
are color-coded to match the corresponding mass in (B). The dynein heavy chain
(blue) binds directly to the dynein intermediate chain (pink) and light intermediate
chain (yellow), while the light chains (green) associate with the intermediate chain
N-terminus. (C) Crystal structure of the dynein AAA+ domain with nucleotide (red)
present in all binding sites 8. (D) Bar diagram of the dynein heavy chain with the
dimerization domain, IC and LIC binding sites, the linker domain and the AAA+
ring domain indicated. (E) Cartoon of the dynein cross-bridge cycle showing the
movement of the linker across the AAA+ ring, allowing for force generation and
stepping.
6 AAA1 is preceded by 543 amino acids, known as the “linker” domain, that connects the motor domains to the dimeric tail (described below). Hydrolysis of
ATP and product release at AAA1 yields a dramatic change in the orientation of the linker with respect to the AAA+ ring, which is accompanied by a conformational change in the linker itself. Prior to ATP binding at AAA1, the linker spans the ring from AAA1 to AAA4 and the microtubule binding domain is in the high-affinity state (Figure 1-1 E). During the process of ATP binding and hydrolysis to ADP-Pi, the linker bends and repositions to the junction of AAA2 and 3 52. Conformational changes within the ring are conveyed to the microtubule binding domain, producing a low-affinity state 56. Release of ADP and Pi allows the linker to return to its original unbent position, which is accompanied by rebinding at a new site on the microtubule (Figure 1-1 E). The large-scale movement generated by the bending of the linker (“power-stroke”) causes the AAA+ ring domain to move along the microtubule and take a “step”.
For dynein to take consecutive steps along the microtubule, two heads need to be coordinated. This occurs through the N-terminus, which acts to dimerize the heavy chain, tethering the two motor and linker domains together into a single unit. This dimerization occurs at the N-terminus of the heavy chain,
AA 1-178 57 (Figure 1-1 B & D). In addition to dimerization, AA 1-1383 of the dynein heavy chain provides two major functions: (1) binding of the IC, LIC and
LCs, and (2) cargo binding. Both the IC and LIC bind directly to the heavy chain, at AA 446-701 and AA 649-800 respectively 58, and the LCs associate with the dynein complex via the N-terminus of the IC, AA 70-150 (Figure 1-1 B & D) 59,60.
7 The dimerized tail with bound IC, LIC and LCs provides a platform for cargo binding, either directly or through the dynein activating complex and cargo adaptor, dynactin in association with coupling scaffolds (discussed later). The tail is also proposed to provide binding sites for other dynein motor regulatory components, such as NudE-Lis1 (see below).
Identification of the “dynein genetic pathway”:
Biochemical purification of cytoplasmic dynein led to its identification and initial characterization 61,62. In parallel, genetic work resulted in the identification of additional proteins and protein complexes that were functionally linked to dynein, the first of which was the S. cerevisiae dynein HC 5,29,35. Targeted mutation of the HC gene led to bi- or multi-nucleated mother cells and anucleate daughter cells, as the plane of nuclear division is within the mother cell (Figure 1-
2 A & B). These mutants were also slow growing, with doubling times increasing by 35-50% at both high and low temperatures. Mutants phenocopying this were identified with random mutagenesis, leading to the discovery of the dynactin components: dynamitin (JNM1), Arp1 (ACT3 or ACT5) and p150Glued (NIP100)
63,64,65{Kahana, 1998 #207,66. A parallel synthetic-lethal screen aimed at identifying genes necessary for viability in the absence of the mitotic kinesin, Cin8, yielded the so-called “perish in the absence of cin8” (PAC) mutants 67. The resulting mitotic associated genes identified additional dynein pathway components, including dynein intermediate chain (PAC11) and Lis1 (PAC1) 67.
8
9 Figure 1-2. Phenotypes of dynein pathway mutants. Dynein pathway mutants
in S. cerevisiae have nuclear localization defects, rather than positioning the
nuclei and spindle within the bud neck to ensure proper division (B), the
spindle is entirely within the mother cell in most mutants (A) 5. The Ropy
phenotype observed in N. crassa is characterized by curled hyphal growth (C),
which markedly differs from the straight growth observed in wild-type N.
crassa (D). Additionally the nuclei fail to migrate into the hyphae in both N.
crassa and A. nidulans mutants (E & G), whereas in wild-type fungi the nuclei
are relatively evenly spaced (F & H) 6,7. Mutation of the p150Glued gene in D.
melanogaster, originally termed the “glued mutant” (I) shows abnormal eye
formation, lacking the hexagonal organization observed in wild-type eyes (J) 9.
10 Soon after the S. cerevisiae HC gene was identified, sequencing of previously discovered nuclear distribution (Nud) and ropy mutants, in A. nidulans and N. crassa respectively, found additional dynein pathway components. Both the Nud and ropy mutants were characterized to have normal nuclear divisions, but the nuclei were not distributed along the length of the growing hyphae (Figure
1-2 E-H) 68,69. The ropy mutants were also characterized by the growth of the hyphae, which are curled, rather than straight, as their name suggests (Figure 1-
2 C & D) 69. Among the proteins identified in these organisms were the dynein
HC (NudA or ro-1), NudE, Lis1 (NudF), LC8 (NudG), and dynactin Arp1 (NudK or ro-4), and p150Glued (NudM or ro-3) 7,70-81, indicating that the “dynein genetic pathway” comprises multiple complexes including dynein, dynactin, NudE and
Lis1.
The dynein motor was confirmed to be essential in metazoans through analysis of the Glued mutant in D. melanogaster, which are homozygous lethal 82.
Sequencing of the Glued gene found the mutation to lie within a novel gene 83, that was soon identified as the homolog of the largest dynactin subunit, p150Glued
84-86. Subsequent identification of the dynein HC gene and mutagenesis resulted in the same eye deformities observed with heterozygous Glued mutants, which lack the normal hexagonal organization (Figure 1-2 I & J) 82. The double Glued,
HC mutants yielded the same phenotype as the single mutants, suggesting that they function together in metazoans, confirming that the pathway is conserved.
Although these screens detected multiple components of the dynein motor, they did not confirm all of the biochemically identified dynactin subunits 3,87,88, as
11 the Arp11, p62, p25 and p27 subunits were missing in these screens. Analysis of ropy mutants not identified as homologs of the dynein motor subunits showed that ro-7 is the Arp11 homolog, but was not initially identified as such because
Arp11 had yet to be discovered 75,87,89. Likewise, this analysis found that the published ro-2 sequence was only a partial sequence, which prevented its identification as p62 89,90. Sequencing of ro-12, which shows the ropy phenotype, but was ignored as it does not have a conspicuous nuclear migration defect, identified it as the p25 homolog. Despite being dispensable for nuclear migration, ro-12 is essential for vesicle trafficking and vacuole distribution within the hyphae
89. Using the ro-12 sequence, the A. nidulans p25 homolog was identified, and mutagenesis also resulted in defective endosomal transport, but not nuclear migration defects 91. Additional analysis found that these vesicle trafficking defects were identical to those seen with the dynein HC mutants, showing that they function within the dynein pathway 89,91.
These screens clearly linked dynein motor function to dynactin and the
NudE-Lis1 complex. Additional confirmation that the genes identified in S. cerevisiae, A. nidulans, N crassa and D. melanogaster are essential in metazoans came from a RNAi screen in C. elegans, in which depletion of dynein motor components was embryonically lethal 92. In this screen, lethality resulted due to defects in pronuclear migrations within the single cell embryo, preventing fusion of the maternal and paternal pronuclei, or spindle defects that prevented cell division. This demonstrated the conserved role of the dynein motor in both nuclear migration and mitosis.
12 Conformational regulation of dynein activity:
External regulators have unambiguously been shown to alter dynein function, but the mechanism by which they do so remains undefined. To think about this, it is important to keep in mind that the motor may be subject to conformational autoinhibition, and that extrinsic binding partners may contribute to activation. A simple model for this is seen with conventional kinesins, which exhibit an autoinhibited state in which the motor domain and cargo binding tail domain interact, essentially folding the molecule in half (Figure 1-3 A) 1. This intramolecular interaction prevents motility by physically blocking microtubule binding by the motor domain. This configuration locks the motor in an ADP- bound state. Cargo binding to the tail results in displacement of the motor domain, which permits microtubule binding. This is thought to promote release of
ADP, with subsequent binding and hydrolysis of ATP coupled to conformational changes that yield motility (Figure 1-3 A) 1.
Dynein also adopts distinct conformations that have been proposed to correlate with activity (Figure 1-3 B-D) 2. When viewed by EM, some particles take on a compact configuration, in which the motor domains appear to be
“stacked” back-to-back, such that the microtubule binding stalks point in opposite directions. These are referred to as Phi particles, due to their resemblance to the
Greek letter ϕ 4. For years, it remained unclear whether the Phi particle represented a physiological conformation, or an EM artifact.
Recent work using recombinant dynein dimers allowed this question to be addressed. In this work, the dynein motor domain and a portion of the linker,
13 14 Figure 1-3. Dynein and kinesin utilize different autoinhibitory mechanisms.
(A) Inhibition of conventional kinesin occurs through an interaction between
the motor domain and cargo-binding domain (left). This interaction is released
through cargo binding, allowing for motility (right) 1,2. Dynein autoinhibition
occurs through AAA+ ring stacking (B-D), generating Phi particles 4. This ring
stacking results in the microtubule binding domains crossing and orienting in
opposite directions, as seen in the EM image (C) and cartoon (D) 2. (E) While
the Phi particles are able to bind microtubules (left), they are not motile;
application of an external force causes the dynein motor to become motile
through rearrangement of the entire structure (right).
15 starting at AA 936, 1004,1110 or 1306, were dimerized with either GST or an extended stiff linker that allowed the physical spacing of the motor domain to be controlled 2. In this system, particles that had a higher propensity to adopt the
Phi configuration as observed with EM exhibited reduced motility in single molecule assays. Physical separation of the AAA+ ring domains yielded increases in both velocity and processivity of movement. All constructs with reduced Phi particle propensity showed increased activity as compared with those that exhibited a high percentage of Phi particles 2.
Understanding how this autoinhibition might be relieved in native motors required a different approach. Attaching two or more motors to the same cargo was also found to yield processive motility, regardless of the propensity for Phi particle formation 2. This suggests that application of an external force to the motor can also relieve autoinhibition 2, leading to a model in which pulling on the tail domain of a microtubule associated motor separates the heads enough to overcome autoinhibition (Figure 1-3 E) 2.
Extrinsic regulation of cytoplasmic dynein:
Extrinsic components of the genetically defined motor might help properly orient the dynein heads, preventing the Phi configuration. Lis1 binds one face of the AAA+ ring domain and has been proposed to function as a “wedge” that separates the motor domains 93-95. Native Lis1 is a dimer, so it has the potential to bind both heads at once. This would be expected to physically constrain the motor domains and prevent head rotation about the linker, preventing Phi particle
16 formation. Because the Lis1 bound motor is not motile, this might represent an intermediate state in which dynein is poised in the correct conformation for movement 93,95. Cargo binding and Lis1 release may provide the final activation step.
Recent EM studies evaluated the configuration of dynein bound to dynactin that include the complex stabilizing protein, BicD2 57,60. The resulting structures show that a kink is imparted into one of the HC N-termini, causing an apparent shift and rotation of the IC β-propeller domains with respect to one another 57. If transduced through the linker domain, this is likely sufficient to separate the AAA+ ring domains, as observed in our microtubule bound supercomplex structure (discussed in Chapter Three). This may account for the increase in motility observed with these super-complexes 96,97.
In summary, the notion that dynein might be subject to structural autoinhibition has only recently begun to be studied. Likewise, understanding how extrinsic regulators alter dynein behavior to allow for both the “stalled” motor activity (dynein plus NudE and Lis1, discussed below) and highly processive motility (stabilized dynein-dynactin super-complex, discussed later in this chapter) is only beginning to emerge. Comprehensive knowledge will require integration of functional and structural data showing how these extrinsic factors alter the dynein motor.
17 The stalled, high-force generating motor complex: dynein, NudE and Lis1:
NudE and Lis1 were first linked to dynein in genetic screens looking for defects in nuclear migration (Figure 1-2) 7,70,75,98. The Lis1 protein is named for the lissencephaly (“smooth brain”) phenotype seen in mammals with mutations in the Lis1 gene 99-103. During normal brain development, neuronal migration and nuclear migrations within neurons stratify the brain cortex, allowing it to increase in thickness. This leads to the formation of the surface folds seen in the brain 100-
104.
Perturbation of the dynein motor also yields defects in mitotic spindle orientation and organization 105-108. Spindle defects can be recapitulated in an in vitro system generated from X. laevis egg extracts, in which dynein, dynactin,
Lis1, NudE or its homolog, NudE-like (NudEL) can be depleted and/or replaced with mutant forms 109,110. In this system it was observed that depletion of NudE(L) prevented focusing of microtubules at spindle poles, which could be rescued by addition of excess exogenous Lis1 (Figure 1-4 A & B). A similar reconstitution experiment in which wild-type NudE(L) was replaced with mutants that could not bind dynein or Lis1 did not result in rescue. Such studies led to the conclusion that NudE(L), which has binding sites for both Lis1 and dynein, and enhances the interaction between Lis1 and dynein, functions as a tether or loading factor that recruits Lis1 (Figure 1-5 A & B) 109,110.
Consistent with this model, formation of the dynein-Lis1 complex with purified components is greatly aided by the addition of wild-type NudE(L) 109,110.
These reconstituted complexes can be assayed using optical trapping, which
18
19
Figure 1-4. Spindle focusing defects observed in the X. laevis egg extract system.
(A) The spectrum of observed spindle focusing phenotypes, the wild-type focused bi-polar spindle (left) to the unfocused “fence” phenotype”. (B)
Histogram showing NudE(L) depletion causes unfocused spindles, and an excess of exogenous Lis1 rescuing the spindle focusing phenotype (modified from 109).
20 21 Figure 1-5. Loading of the dynein-NudE(L)-Lis1 complex. (A) NudE(L) (yellow)
binds to the dynein tail through direct interactions with the dynein IC N-
terminus (pink, arrowhead 1) and LC8 (green, arrowhead 2). These
interactions generate a stable complex that is able to tether Lis1 (red,
arrowhead 3) to the dynein motor complex. This tethering increases the
likelihood of Lis1 binding to the AAA+ ring of dynein (B) generating a stalled
motor complex. Cartoon is drawn to scale.
22 demonstrated Lis1 (or Lis1 and NudE(L)) caused dynein coated beads to bind microtubules persistently and allows the motor to withstand significantly greater pulling force before detaching as compared to beads coated with dynein alone 111.
Persistent binding of the dynein-Lis1 complex continued in the face of ATP hydrolysis, indicating that in the presence of Lis1, ATP hydrolysis at AAA1 fails to result in the conformational changes required for motility, essentially “stalling” the dynein motor 93,94. Taken together, these findings have led to a model wherein
Lis1 acts as a molecular clutch, decoupling dynein stepping from ATP hydrolysis, to allow the motor to remain microtubule bound while tolerating high-load forces.
My work investigating the mechanism by which the “stalled” dynein-Lis1 complex is formed will be presented in Chapter Two.
Discovery and structure of the dynein-activating complex, dynactin:
The Dynein-NudE(L)-Lis1 complex inhibits dynein movement, whereas dynactin has the opposite function, activating dynein 12,84. The components of the 1.2 MDa dynactin complex were identified through the combination of genetic screens (discussed previously) and biochemistry. Electron microscopy studies indicated that dynactin is organized into three structural domains 112. The core of the dynactin particle is a short actin-like filament, that is associated with a shoulder/arm domain at one end and small protruding complex at the other
(Figure 1-5 A & B) 112. Unlike the homodimeric dynein complex, dynactin’s eleven subunits are present in varying stoichiometric ratios and organized into a highly asymmetric structure 3,87,88.
23
24 Figure 1-5. Dynactin composition and subunit organization. (A) Coomassie
Blue stained SDS-Polyacrylamide gel of purified bovine dynactin. Subunit
labels are color-coded to match the corresponding mass in (B). (B) Cartoon of
the dynactin structure with the subcomplexes and subunits indicated. The
shoulder/arm has been shown to interact with the dynein tail, whereas the Arp
filament and pointed end complex are responsible for cargo binding.
25 Despite the structural asymmetry of the dynactin molecule overall, the shoulder/arm domain (orange and yellow in Figure 1-5), which is the subject of
Chapter Four and Appendix II, has a dimeric composition and appears to exhibit mirror symmetry when viewed in EM 57,60. It contains two protomers of p150Glued and p24, and four protomers of dynamitin/p50 (DM) (Figure 1-5 A & B) 87. The projecting arm, which consists of the N-terminal ~1050 amino acids of p150Glued, accounting for ≈ 117 kDa, or 82% of its mass 57. It binds dynein AA 1-32 in a four-helix bundle with p150Glued AA 415-530 113,114. The N-terminal domain of p150Glued, which appears as a small globular domain in the EM, contains a CAP-
Gly motif and basic domain, AA 25-105 and AA 115-145 respectively, both of which can bind microtubules directly 115. Given the importance of p150Glued to dynactin functions 113,115-120, gaining an understanding of how it is incorporated into dynactin is clearly worthwhile.
How the shoulder/arm complex attaches to the Arp filament is known.
Dynamitin (AA 1-88) binds Arp1 directly, with each dynamitin associating with two Arp1 protomers 57,121. This interaction anchors the entire shoulder/arm complex to the Arp filament, allowing the dynein and microtubule binding activities of dynactin to be structurally coupled to the domain of dynactin that interacts with cargo.
Dynactin complex structure has largely been solved using EM, and fitting available crystal structures and homology models into the EM envelope 57,60,122;
(discussed in more detail in Chapter Three). Despite the new insights EM has yielded on the structure of the Arp filament, the shoulder domain remains
26 mysterious. This is because the resolution obtained for the shoulder in EM models was somewhat lower than for the Arp filament (4.0 Å vs. 6.3 Å) 57, and the fact that no homology models are available for dynamitin and p24, making model fitting impossible. Despite these limitations, the shoulder domain was found to be a symmetric structure composed primarily of α-helices that form two
U-shaped three-helix bundles. Because the length of each arm of the U-shape is different, helices predicted from the p24 and dynamitin sequences can be ascribed to different parts of the three-helix bundle. This model and my biochemical work verifying it is the topic of Chapter Four.
Stabilization of the ultra-processive dynein-dynactin-scaffold complex:
In vitro microtubule gliding or bead motility assays have shown that dynein alone is able to move cargo. Dynactin does not alter the velocity of dynein-based vesicle or bead movement, but rather acts as a mild processivity enhancer 12,84.
By contrast, single fluorescently labeled dynein particles do not undergo processive movement even in the presence of dynactin 96,97. Together these findings suggest that dynein may exist in multiple states that are not fully active for transport: (1) an autoinhibited dynein state, as previously described (Phi particles), that can bind microtubules but not move, (2) a Lis1 inhibited state that binds microtubules and withstand high pulling forces, and (3) a dynactin bound state that is weakly processive and needs additional regulation to yield highly processive motility.
27 The protein Bicaudal D2 (BicD2) has emerged as the archetype of the third type of regulator, which acts a scaffold to facilitate interactions between dynein and dynactin. The D. melanogaster ortholog, Bicaudal, is required for dynein based mRNA transport in oocytes 25,27. Studies in cultured mammalian cells indicate that BicD2 is predominantly associated with the Golgi complex.
Overexpression of N-terminal fragments induce steady state localization changes of the Golgi complex, similar to the alterations observed with other dynein perturbations, and is likely due to interactions among dynein, dynactin and exogenous BicD2 fragments that block interactions with the Golgi complex associated endogenous BicD2 123,124. Supporting this phenotypic observation, dynein and dynactin components co-immunoprecipitate with the BicD N-terminus
123. Together these findings suggest that BicD2 interacts with dynein and dynactin in vivo.
Follow up in vitro studies showed that a fragment of BicD2 (AA 25-400) was able to cause purified dynein and dynactin to sediment more rapidly in sucrose density gradients, suggesting super-complex formation 125. Final proof that BicD2 plays an activating role in dynein-based motility was obtained via single molecule assays of fluorescent complexes. The BicD2 fragment was found to act as a scaffold that promotes formation of an ultra-processive dynein complex that moved over very long distances, with no change in the velocity 96,97.
The structure of the dynein-dynactin-BicD2 (DDB) complexes, bound to microtubules will be discussed further in Chapter Three.
28 Other proteins, including Spindly, Hook3, and Fip3 were found to stabilize the dynein-dynactin associating and yield similar ultra-processive complexes 96.
These proteins all have an extended coiled-coil domain, suggesting a common mode of binding, and contain another domain that is proposed to interact with cargo, either directly or indirectly. Another common theme is that these proteins bind specific Rabs, suggesting a mode of regulation. Further studies are necessary to mechanistically tease apart the role each of these scaffolds play in dynein-based movement.
Despite the many recent advances towards understanding structure and function of the dynein motor, many questions still remain. It is clear that
NudE(L)-Lis1 and dynactin promote different dynein functions, but how all the components of the motor complex are integrated in vivo remains mysterious.
The work presented in the following chapters provides novel insights into the complicated dynein motor, both at the single molecule level and in bulk biochemical studies. These studies provide a better understanding of dynein motor regulation, which will provide a strong basis for future work.
29
Chapter Two
Examining the loading mechanism of the stalled dynein-
NudE(L)-Lis1 motor complex
30 Introduction:
The dynein motor transports cargo along microtubules, from the plus end towards the minus end 21-23. These cargoes can be divided into “low-“ and “high- load”, based upon the amount of force and number of motors required for transport. Low-load cargoes include vesicles, mitochondria and other organelles
12,126-129; whereas, “high-load” transport includes nuclear movement and mitotic spindle organization 130,131 5 21 35 70.
Screens identifying mutants with spindle or nuclear positioning or migration defects revealed the components of the dynein motor (discussed in
Chapter One). The components identified were the dynein complex itself and the adaptors dynactin, NudE and Lis1, which alter dynein motility 5,6,25,29,30,63-65,67-
69,71,73,76-81,83,89-91,132-144. These screens also demonstrated that the dynein motor is essential. In budding yeast, positioning of the nucleus and mitotic spindle to the bud neck ensures both daughter cells receive a full complement of genetic material 35,145. Nuclear positioning and migration is also crucial in single and multi-nucleated cells, such as neurons and the hyphae of filamentous fungi. In neurons nuclear migration helps to stratify developing tissues, such as the brain.
Without these nuclear translocations, the developing tissues are improperly innervated, and generally have loss of function 101,102,146,147. Unlike neurons, the hyphae, or germ tube, of filamentous fungi are multi-nucleated, with the nuclei spaced evenly throughout the length of the hyphae. Fungi lacking the required dynein motor components are unable to transport the nuclei into the hyphae, thus preventing spore production 7,75,98,100,129,148.
31 More recently, it has been shown that mutations within the dynein motor can lead to improper division of stem cells as well. In these cells, spindle positioning determines whether division of the progenitor stem cell is symmetric, yielding two identical progenitor or differentiating cells, or asymmetric, with the division producing one progenitor cell and a differentiating cell. Altering the balance between symmetric and asymmetric divisions can have drastic consequences, including depletion of stem cells or decreased numbers of differentiating cells. Both of these consequences have been observed in hematopoietic stem cells, D. melanogaster germ-line and neural stem cells, where they are associated with various disease states 106,107 ,149-151.
These mutational studies indicated that the processes of nuclear and mitotic spindle positioning and nuclear migration were dependent on dynein, dynactin, Lis1 and NudE, or its homolog NudE-Like (NudEL). However, it remained unclear how NudE(L) and Lis1 were affecting dynein functions. More specifically, the studies did not address whether these two proteins interact and directly regulate the dynein motor, or if they act as indirect regulators. Cultured neurons and fibroblast cells were utilized in attempts to more clearly define this regulation, where it was noted that dynein and Lis1 had similar subcellular localizations. Furthermore, experiments in cells over-expressing tagged Lis1,
NudE(L) and/or the dynein intermediate chain (IC) indicated that these proteins might interact, as they exhibited similar alterations in localization patterns. The interactions among these three proteins were further supported by coimmunoprecipitation data, but whether it was through direct or indirect binding
32 could not be determined 152,153. Yeast two-hybrid and co-expression studies using fragments of dynein HC and IC, NudE(L) and Lis1 first showed that the interactions among these components were direct. In this work, NudE(L) appeared to bind both dynein HC and IC, while Lis1 was found to only interact with the HC. Previously observed interactions between NudE(L) and Lis1 were confirmed in this work as well 152,153.
Refinement of these findings used recombinant full-length proteins and protein fragments in vitro. These studies indicated that NudE(L) binds the IC N- terminus and helps Lis1 interact with the AAA+ domain of dynein, blocking the linker swing. This results in a stalled motor, despite continued ATP hydrolysis
93,94,111. Much of the work determining the role of Lis1 was performed with recombinant dynein heads. Although it was possible to observe Lis1-dynein binding, it required concentrations of Lis1 that were much higher than physiological conditions, suggesting that in vivo the presence of NudE(L) in vivo may help facilitate this interaction.
Support for this idea came from studies in Xenopus egg extracts, where the absence of Lis1, NudE(L), dynein or dynactin all result in aberrant spindle formation. Depletion of any of these components yields spindle microtubules that are not focused at the centrosomes. Rather, they display a range of phenotypes from multipolar spindles, to a complete failure to focus microtubules, or a fence phenotype (Figure 2-1 A) 110. However, it was possible to rescue the spindle aberrations caused by depletion of NudE(L) by adding exogenous Lis1, such that
33
34 Figure 2-1. The dynein motor is required for proper spindle formation in vitro (A)
Range of spindle defects observed with dynein motor component depletion in
Xenopus egg extracts (from reference 110). Scale bar is 10 µm.
35 the Lis1 concentration exceeded that normally observed in vivo. Furthermore, adding exogenous NudE(L) or an N-terminal fragment, AA 1-201, was sufficient to rescue the spindle abnormalities. By contrast, NudE(L) fragments that were unable bind to either the dynein IC or Lis1 could not rescue the phenotype
(Figure 2-2 A) 110.
The observed phenotypic rescue yielded a testable loading model, where
NudE(L) acts as a tether between dynein and Lis1. This tethering function occurs via the N-terminal 80 amino acids of NudE(L), which interacts with the dynein IC, and AA 100-153, which bind the N-terminus of Lis1 109. The additional interaction between NudE(L) and dynein physically links Lis1 to dynein, helping it to bind to the AAA+ domain and generate a stalled dynein motor complex 111.
Testing this loading model in vitro requires the use of holo-dynein complexes, rather than the recombinant AAA+ head domain used in the previous experiments. Our lab has successfully purified active dynein motors from chick embryo and bovine brain, both of which were used to examine the direct interactions among dynein and recombinant NudE(L) and Lis1 (Figure 2-2 A & B).
Although we cannot exclude the possibility of other NudE(L)-dynein subunit interactions, my analysis with a recombinant dynein IC fragment (Figure 2-2 C) confirmed NudE(L) binds dynein via a direct interaction with this subunit.
36
37 Figure 2-2. Recombinant protein fragments. Schematic showing the domains of
(A) NudEL, (B) Lis1 and (C) DIC. The boxed regions indicated on (A) NudEL and
(C) DIC are binding domains for the protein named therein.
38 Results:
NudE(L) fragments interact differentially with the dynein holo-complex:
To investigate the dynein-NudE(L) complex, I took advantage of the large
S-value of dynein(≈20 S), as most proteins exhibit smaller sedimentation values
(Figure 2-3 A). NudEL, an extended coiled-coil protein, sediments around 4 S
(Figure 2-3 B), as do the smaller NudEL fragments (Figure 2-3 D & E). It has been established that this behavior can be altered by complex formation.
Therefore, I looked for NudEL to shift into the ≈20 S fractions with dynein to determine if direct binding occurred.
The first experiment performed utilized full-length recombinant NudEL, ensuring that all dynein binding domains were present. When combined with dynein, then subjected to sedimentation, the behavior of NudEL is altered. Some of the protein still sedimented as a small particle, in the light sucrose fractions.
Additionally, the NudEL peak broadened, with NudEL sedimenting in the ≈20 S fraction with dynein, indicating complex formation (Figure 2-3 C). Interpretation of this result was complicated by the observed degradation of the NudEL protein.
These degradation products showed differential interactions with dynein, with the larger degradation products sedimenting ≈20 S, while smaller fragments did not interact.
Given that the full-length NudEL protein degrades, I set out to determine whether shorter, more stable fragments might mimic the behavior of the full- length protein. Previous studies used two different NudEL fragments, either
39
40 Figure 2-3. Dynein forms a stable complex with the coiled-coil domain of NudEL.
(A) The dynein complex sediments as a large 20 S particle. (B) Full-length and fragments of NudEL, AA 1-201 (D) and AA 8-171 (F) sediment as small particles at the top of the sucrose density gradient. Both Full-length NudEL (C) and AA 1-
201 (E) shift into the ≈20 S fraction in the presence of dynein. (G) The behavior of NudEL AA 8-171 is unaltered by dynein. Arrows indicate the position of sedimentation standards.
41 amino acid 1-201 or 8-171 109,110,154. Both of these fragments contain the dynein
IC binding site; however, only the longer fragment contains a second reported dynein binding site for LC8 155. The longer fragment, AA 1-201, appears to bind dynein, as evidenced by cosedimentation in the 20 S pool (Figure 2-3 E). The shorter NudEL fragment, AA 8-171, did not have an altered behavior in the presence of dynein (Figure 2-3 G), indicating that the LC8 binding site may be necessary to form a stable dynein-NudEL complex in solution.
Dynein IC interacts with NudE(L) fragments, but not full-length NudE(L):
The sedimentation studies suggest that the NudEL-dynein interaction does not rely entirely on the interaction between NudEL and the IC. Therefore, I set out to further investigate potential interactions using recombinant fragments of dynein IC and LC8 in combination with different NudEL fragments. Here I used size exclusion chromatography (SEC) to investigate the interactions between these subunits, as had been performed for the IC-p150Glued interactions
113. I first determined how individual fragments behaved when subjected to SEC.
This data will allow me to detect shifts in the elution profile when protein mixtures are analyzed, which are indicative of complex formation.
SEC analysis of the dynein IC fragment, AA 1-106, indicated that it behaved as a small particle with a Stokes’ radius of 32.9 ± 1.6 Å (Figure 2-4 A).
Analysis of full-length NudEL and the shorter fragments show that they all behave similarly and appear larger than IC AA 1-106. Full-length NudEL has a
Stokes’ radius of 55.4 ± 1.2 Å (Figure 2-4 B), which is identical to the Stokes’
42 43 Figure 2-4. DIC interacts with NudEL fragments but not full-length NudEL.
Superdex 200 analysis (left) and Coomassie Blue stained SDS-PAGE (right) showing included column volume. Analysis was performed for DIC (A), full-length
NudEL (B) and the NudEL fragments AA 1-201 (D) and AA 8-171 (F) alone.
Incubation of DIC with different NudEL fragments show a shift in the elution behavior of DIC when combined with AA 1-201 (E) and AA 8-171 (G), but not full- length NudEL (C).
44 radius of 54.9 ± 0.12 Å observed for the NudEL fragment, AA 1-201 (Figure 2-4
D). This similarity suggests that models wherein the N-terminal coiled-coil domain of NudEL interacts with its C-terminus may be correct. This fold-back model would explain why full-length NudEL and the fragment appear to have similar dimensions in solution. Unlike full-length and AA 1-201, the shortest
NudEL fragment, AA 8-171, eluted as a broad double peak, with Stokes’ radii of 54.4 ± 0.63 Å and 49.3 ± 0.38 Å (Figure 2-4 F). As the
NudEL and IC fragments could be resolved with SEC, it was possible to determine whether these proteins bind each other in solution.
To our surprise, full-length NudEL was unable to bind to the dynein IC fragment (Figure 2-4 C). Incubating NudEL with the dynein IC showed identical elution behavior as when the proteins were analyzed alone. However, the shorter NudEL fragments were able to bind to the IC fragment, with the majority of the IC AA 1-106 shifting to elute earlier with NudEL fragments. In both instances, the NudEL fragment-IC complexes behaved similarly to the NudEL fragment alone; with the AA 1-201-IC complex eluting with a Stokes’ radius of
55.1 ± 0.07 Å and the AA 8-171-IC complex exhibiting a double peak with
Stokes’ radii of 53.2 ± 0.86 Å and 50.3 ± 0.45 Å (Figure 2-4 E & G). Combining the size information obtained for full-length NudEL with its inability to bind the dynein IC, under the same conditions that interactions were observed with shorter NudEL fragments, further supports the model that NudEL undergoes intramolecular interactions in solution that may affect dynein binding.
45 After investigating the IC-NudEL interactions, I examined the interaction between NudEL and LC8; however, these results were not reproducible. The interactions between NudEL and LC8 appeared to be somewhat weak (data not shown). The most promising result showed a small amount of LC8 coeluting with the NudEL fragments, as compared with the drastic shift observed with the IC fragment. Increasing protein concentrations had little impact on the reproducibility of these results. NudEL-LC8 affinity measurements have not been published, suggesting that this interaction may be too weak to measure.
The direct NudEL-LC8 interaction was initially detected via pull down assays with recombinant protein fragments 155. However, exogenous LC8 was not able to competitively inhibit the NudEL-dynein interaction, even though exogenous IC was able to block NudEL-dynein complex formation in the same assay 155. This suggests that the NudEL-IC interaction brings NudEL into the proximity of LC8, increasing the likelihood of an interaction between NudEL and
LC8 that further stabilizes the dynein-NudEL complex. However, this LC8-
NudEL interaction is likely low affinity, and unstable in solution, explaining the difficulty in generating reproducible binding results.
Lis1 requires NudEL to bind the dynein holo-complex:
Although the previous experiments demonstrated that a stable dynein-
NudEL complex can form, such a complex is not a stalled, high-force generating motor. The stalled dynein motor requires binding of Lis1 to the AAA+ ring domain
93,94,111. Therefore, I next probed the dynein-Lis1 interaction with the
46 47 Figure 2-5. Dynein-Lis1-NudEL form a ternary complex. (A) Dynein sediments as a 20 S particle when subjected to sucrose density gradient analysis. (B) Lis1 sediments as a small particle at the top of a sucrose density gradient. (C) When combined with dynein, the behavior of Lis1 is unchanged. (D) Lis1 and NudEL cosediments in the light fractions of a sucrose density gradient, but when combined with dynein, they cosediment as a 20 S complex (E). Brackets in B-D show the peak broadening observed with Lis1. Arrows indicate the position of sedimentation standards.
48 cosedimentation assay that I previously used to examine dynein and NudEL binding (Figure 2-3). In this assay, I found that recombinant Lis1 sediments in the lighter sucrose fractions, ≈5 S (Figure 2-5 B). Contrary to published results, there was no indication that Lis1 cosedimented with the dynein complex (Figure
2-5 C). This may be a result of the nucleotide state of the dynein AAA+ domain, as the dynein had been cryo-stored in an excess of ATP (2 mM), and Lis1 was shown only to bind recombinant mammalian dynein heads when incubated with
ATP- Vanadate, which mimics the ADP-Pi bound state. By contrast, incubation of the recombinant mammalian dynein head with either ATP or ADP did not yield a complex 111. This differs from studies with yeast dynein heads, where nucleotide state had no impact on Lis1 binding 93.
Although I was unable to generate a complex with only dynein and Lis1, it seemed likely that the presence of NudEL may enhance complex formation, as has been shown in vitro with Xenopus egg extracts 109-110. To examine this, I first confirmed that the Lis1-NudEL complex could be resolved from a ≈20 S complex.
This analysis indicated that the Lis1-NudEL complex sediments as a broader peak than either protein alone, but it is still ≈5 S (Figure 2-5 D, bracket). By incubating dynein, Lis1 and NudEL and subjecting the resulting mixture to sucrose density gradient sedimentation, it was apparent that a complex forms.
Both Lis1 and NudEL exhibit broader sedimentation behavior, with both appearing in the ≈20 S fractions of the gradient (Figure 2-5 E). This supports the model that NudEL functions as a loading tether for the dynein-Lis1 complex both in vitro and in vivo.
49 Discussion:
The data presented here appear to be internally conflicting. Full-length
NudEL binds the dynein holo-enzyme (Figure 2-3 B & C), but a direct interaction with the dynein IC fragment cannot be detected (Figure 2-4 B & C). Conversely, both fragments of NudEL are able to bind directly to the IC fragment in solution
(Figure 2-4 D-G), but show differential binding to holo-dynein (Figure 2-3 D-G).
These apparently contradictory results may be indicative of more complicated interactions between NudEL and dynein. The simplest explanation would be an intramolecular interaction that occurs with the full-length NudEL protein, but not the fragment. Releasing this intramolecular interaction may be dependent on binding to multiple dynein subunits and/or Lis1, as suggested by the different binding behavior of the NudEL proteins with the dynein IC and the dynein holo- complex.
In this model the direct interaction between NudEL and the IC is required for complex formation, but the behavior of the shortest NudEL fragment suggests that this interaction is not sufficient to generate a stable complex. My attempts to confirm that the reported direct interaction between LC8 and NudEL may be responsible for stabilizing this were inconclusive. Going forward, point mutations within the reported LC8 binding site, NudE AA 192-211, could be used establish the importance of LC8-NudEL binding 111. This mutant could be analyzed with the dynein holo-complex cosedimentation assay to determine whether these residues are required for stable association of NudEL with dynein.
50 The similar behavior of full-length NudEL and the shorter fragments in the
SEC analysis suggests that in solution these particles have nearly identical dimensions. This supports the models where NudEL interacts with itself, which would block the binding sites for the IC and other binding partners. However, this raises questions regarding how these intramolecular interactions are released to allow NudEL to bind. The methods used here were unable to detect how this may occur, thus, necessitating more work to determine whether release of this inhibition is spontaneous or dependent upon interactions with Lis1 or dynein subunits.
Although I was not able to fully resolve the NudEL-dynein binding mechanism, I demonstrated that the Lis1-dynein interaction is weak, and may depend on the nucleotide state in AAA1 or AAA3. This needs to be explored further. However, Lis1 and NudEL interact to form a dynein-binding complex, which effectively loads Lis1 onto the dynein motor domain. My findings fully support the model of NudEL acting as a tether to load Lis1 110,111,154. Here I expand upon this, showing that the entire coiled-coil domain of NudEL, AA 1-201, is necessary for stable association with dynein (Figure 2-3) and loading of Lis1 onto the dynein AAA+ domain (Figure 2-5).
The work presented here is consistent with current models for the formation of the stalled dynein-NudEL-Lis1 complex, however, little is known about how the dynein motor can switch from a stalled state, to a processive motor. Potentially, this switch may occur via competitive binding between dynactin and NudEL, both of which bind to overlapping sites on the IC 109,113,114.
51 This suggests a mutually exclusive binding of Lis1-NudEL or dynactin, resulting in either a stalled or processive motor complex, presenting a model where the higher affinity p150Glued-IC interaction replaces the NudEL-IC interaction 156,157.
Release of NudEL from the IC may destabilize the binding of the Lis1-NudEL complex, allowing for the formation of the processive dynein-dynactin motor.
This model presents multiple testable hypotheses. First, the assumption that NudEL and p150Glued binding are mutually exclusive. Although competitive binding experiments have been performed, they utilized IC fragments rather than the dynein holo-enzyme 111,156,157. As NudEL appears to require additional interactions with dynein, the binding of NudEL or p150Glued with only the IC may not be representative of it’s binding to dynein. Secondly, release of NudEL may not be sufficient to destabilize Lis1 binding; the role that NudEL plays after it loads Lis1 onto the dynein complex remains unclear. Resolving these questions may present a more complete model of how switching from a stalled to processive motor complex occurs. Fully determining this regulatory mechanism will require a combination of interaction studies and motility assays, using both wild type and binding mutants of all proteins involved.
Materials and methods:
Sf9 expression and purification of Lis1:
His-tagged Xenopus Lis1 was produced in Sf9 cells, as described previously 110. Briefly, Sf9 cells were infected with the His-Lis1 bacculovirus for
72 hours. After infection, cells were pelleted and either lysed immediately for
52 protein purification, or stored as a cell pellet at -20° C for future use. Cell pellets were resuspended in 50 mM Tris, 500 mM NaCl with 1% NP-40, pH 8.0, then dounce homogenized on ice to lyse. After lysis, the supernatant and insoluble cell debris were separated via centrifugation. The supernatant was syringe filtered with a 0.2 µm filter, then loaded onto a 5 ml HiTrap TALON crude column
(GE life sciences). The column was washed with 50 mM NaPO4, 500 mM NaCl,
50 mM Imidazole, pH 8.0 until the baseline returned to zero. The bound protein was eluted using an Imidazole bump to 500 mM Imidazole in 50 mM NaPO4, 500 mM NaCl. Purified protein was analyzed via Coomassie blue stained SDS-
PAGE for purity, and used within 48 hours of purification for binding experiments.
We found that the purified Lis1 protein could not be cryo-stored and that after 48 hours on ice, it had degraded significantly.
Expression and purification of NudEL, p150Glued and DIC fragments:
Full length His-tagged NudEL and fragments were expressed and purified as previously described 110. Bacteria carrying the plasmid to express full-length or fragments of NudEL were grown at 37° C to an OD600 of 0.4-0.6, then induced with 0.1 mM IPTG overnight, 16-20 hours, at 22°C. After centrifugation, the cell pellet was resuspended in 50 mM Sodium Phosphate, 300 mM NaCl, pH 7.4 with protease inhibitors, lysed using an Avestin EmulsiFlex C3. The syringe filtered supernatant was loaded onto a 5 ml HisTrap column (GE Life Sciences), washed until the baseline returned to zero, then eluted with 50 mM Sodium Phosphate,
53 300 mM NaCl, 500 mM Imidazole, pH 7.4. Peak fractions were analyzed with
Coomassie blue stained SDS-PAGE for purity.
His-tagged DIC AA 1-106 was expressed and purified as previously described 113. Briefly, Bacteria carrying the plasmid to express the DIC fragment were grown at 37° C to an OD600 of 0.4-0.6, then induced with 0.1 mM IPTG overnight, 16-20 hours, at 37°C. After centrifugation, the cell pellet was resuspended in 50 mM Sodium Phosphate, 300 mM NaCl, pH 7.4 with protease inhibitors, then lysed using an Avestin EmulsiFlex C3. The syringe filtered supernatant was loaded onto a 5 ml HisTrap column (GE Life Sciences), washed until the baseline returned to zero, then eluted with 50 mM Sodium Phosphate,
300 mM NaCl, 500 mM Imidazole, pH 7.4. Peak fractions were analyzed with
Coomassie blue stained SDS-PAGE for purity.
Dynein purification from bovine brain:
Dynein and dynactin were purified from bovine brain as described previously (Jim Bingham) with the following changes. Dynein and dynactin were eluted from the MonoQ 10/100 column using a linear salt gradient. Dynein and dynactin containing fractions were identified by Coomassie blue stained SDS polyacrylamide gels, then rechromatographed separately on a MonoQ 5/50 column. Elution from the MonoQ 5/50 column was performed with a salt bump, the concentration of salt was determined from the conductivity measurements of the first and last fractions pooled from the MonoQ 10/110 column (described in more detail in Chapter 3).
54 Dynein cosedimentation assay:
50 µg of purified bovine dynein was incubated with combinations of Lis1,
NudEL and/or CC1 protein fragments, in a 1:20 molar ratio of dynein to binding partner. The proteins were brought to a final volume of either 500 µl (SW41 gradients) or 250 µl (SW55 gradients) in 20 mM Tris, 100 mM NaCl, 1 mM DTT, pH 7.2 and incubated on ice for 30 min. After incubation, the protein samples were loaded onto 5-20% sucrose (w/v) gradients in 20 mM Tris, 100 mM NaCl, 1 mM DTT, pH 7.2, then centrifuged at 30,000 RPM for 15 hours in an SW41 rotor or 55,000 RPM for 3.5 hours in an SW55 rotor at 4°C.
Gradients were fractioned into either 14 x 0.9 ml fractions for SW41 gradients or 12 x 0.5 ml for SW55 gradients. 10 µl of each fraction were loaded and run with SDS-PAGE, then transferred to PVDF membrane for immunoblot analysis. Dynein subunits were detected with antibodies against the heavy chain, mAb 440.1, and intermediate chain, mAb 74.1 (Millipore). Full length NudEL or fragments were detected with a polyclonal antibody generated against full length
Xenopus NudEL (gift from Yixian Zheng’s lab) 158. Lis1 was detected using the mAb Lis1-388 (Sigma). Coomassie blue stained SDS-PAGE analysis was run in parallel to the immunoblot analysis. However as several of the proteins have molecular weights of ~45-55 kDa, determining the identity of proteins in this region of the gel with Coomassie blue staining was insufficient.
55 Size exclusion chromatography analysis of protein-protein interactions:
Full length NudEL or fragments were combined with DIC AA 1-106 in a 1:1 molar ratio in 20 mM Tris, 150 mM NaCl, 1 mM β-ME, pH 7.4. Proteins were loaded onto a pre-equilibrated Superdex200 10/300 GL (GE Life Sciences) in 20 mM Tris, 150 mM NaCl, 1 mM β-ME, pH 7.4. A 0.5 ml protein sample was injected into the column at 0.25 ml/min, the column continued to run at 0.25 ml/min with 0.5 ml fractions collected throughout the entirety of the column run.
Stokes’ radii were calculated using a standard curve generated with standards from both the Gel Filtration HMW and LMW Calibration kits (GE Life Sciences) according to the protocol for the Superdex200 10/300 GL column. Peak fractions from the Superdex200 10/300 GL column were analyzed with Coomassie Blue stained SDS-PAGE to determine whether shifts in protein elution patterns were evident.
56
Chapter Three
EM analysis of the dynein, dynactin and dynein-dynactin-
Bicaudal D2 complexes.
57 Introduction:
Dynein and dynactin are both large multi-subunit complexes, with masses of 1.5 MDa and 1.2 MDa, respectively. Although these complexes were identified in 1984 and 1991 12,61,159, high-resolution structures have been slow to emerge, and are limited to the motor domain of dynein. The first motor domain structures solved showed that the microtubule-binding stalk contained an anti- parallel coiled-coil domain. During ATP hydrolysis within the AAA+ ring, conformational changes are transferred into this coiled-coil domain causing the
α-helices to slide relative to one another, which alters the structure of the microtubule-binding domain (MTBD). The distinct structures of the MTBD are assumed to correspond with the high- and low-affinity microtubule-binding states.
Soon after, a recombinant AAA+ ring domain was expressed in Sf9 cells, leading to near atomic resolution crystal structures 8,53. This structural work, combined with electron microscopy and biochemical analysis, has provided great insight into the hydrolytic cycle and how it is coupled to conformational changes driving motility 48,51,54,56. However, this analysis was limited to truncated heavy chain fragments lacking tail domain, limiting insight into the organization of the other dynein subunits.
Low-resolution electron microscopy analysis of both native dynein and dynactin has yielded considerable insight into their structures 4,41,87,112,122,160,161.
The prominent feature observed using either negative stained or platinum coated freeze-etched EM dynein samples is the motor domain. These appear as dimpled rings, which are easily observed due to their high contrast in electron
58 micrographs. It is also possible to make out a V-shaped tail domain that connects the two AAA+ ring domains in some particles.
In these micrographs, the dynein holo-complex exhibits multiple conformations, with the center-to-center spacing of the motor domains ranging from stacked (the so-called “Phi particle”, discussed in Chapter One), to >35 nm apart. It has been speculated that the linker domain, which contains 543 amino acids that connect the AAA+ ring domain to the tail, contributes to this heterogeneity. Conformational and positional changes within the linker are known to be responsible for the “power-stroke”, which is triggered by ATP hydrolysis and product release at AAA1 (as discussed in Chapter One). The importance of the linker to the power-stroke has been well studied, but structural work on this domain was limited until a recent crystallographic study elucidated the “sweep” of the linker across the face of the motor domain 183. This limitation is the result of particle averaging, which can only be used for structures that demonstrate consistent conformation and position. For this reason, using EM to obtain structural information for the linker domain and better definition of the tail domain requires a novel strategy, focused classification, which will be discussed later in this chapter.
EM analysis has also provided a foundational understanding of dynactin structure 87,112,122,160,162. Antibody decoration of single particles yielded an overview of subunit organization within the complex, which was further mapped with biochemical cross-linking, revealing interactions among subunits of the pointed-end complex 163. A 34 Å resolution, 3D averaged dynactin structure
59 yielded additional insight into the overall shape of the particle 122. Fitting of crystal structures and homology models of the CapZ heterodimer and actin- related proteins allowed the maximal number of Arp protomers that could fit to be limited to nine, but this number could not be definitively determined owing to the relatively low resolution of the structure. Furthermore, the presence and location of the conventional actin monomer could not be deduced, again due to resolution limitations. The core structure of dynactin showed very little flexibility or variation, with a consistent length of 34 nm. By contrast, the projecting arm was observed to have a high degree of flexibility, making it impossible to obtain structural information from averaged images. Finally, the organization of the shoulder domain remained unclear, as identification of the α-helices and β-strands that are predicted to comprise this structure is impossible at 34 Å resolution.
Although previous structural work has yielded important insights into dynein and dynactin structures individually, it provided no information about how the two complexes come together to generate the functional super-complex.
Biochemically, a direct interaction between p150Glued and dynein intermediate chain is well established 113,117,118. Agents predicted to block this interaction inhibit dynein-dynactin functions in vivo 113,131,164-167. However, the precise structural locations of the portions of dynein and dynactin responsible for this interaction (p150Glued AA 415-530 and DIC AA 1-32) are unknown 114.
Uncertainties again arose because both the dynactin arm and dynein tail exhibit flexibility, which impedes structural analysis.
60 With only a single, small interaction site identified, a wide range of super- complex structures can be imagined, making modeling of the super-complex difficult. Furthermore, efforts to obtain structural insight into the dynein-dynactin super-complex were thwarted by the fact that dynein and dynactin do not bind each other with high affinity. Despite these problems, it has been the goal of much work to determine a dynein-dynactin super-complex structure. An understanding of this structure is likely to yield considerable information into dynein’s functions and regulatory mechanisms.
In this chapter, I detail my efforts to obtain dynein and dynactin samples suitable for high-resolution frozen-hydrated (cryo) EM analysis. I will also discuss the insights we obtained into the holo-dynein and dynactin structures, as well as the structure of the dynein-dynactin super-complex (stabilized with BicD2) bound to microtubules. These structural studies were performed in collaboration with Gabriel Lander and Saikat Chowdhury in the Department of Integrative and
Computational Biology at the Scripps Research Institute (La Jolla, CA).
Results:
Preparation of chick embryo and bovine brain dynein and dynactin:
Previous EM collaborations used chick embryo brain (CEB) dynein and dynactin prepared in the Schroer lab with great success 87,112,122,160. In contrast, porcine brain dynactin isolated by our collaborator, Hiroshi Imai, yielded heterogeneous particles that were unsuitable for image averaging. Owing to concerns that bovine protein would behave similarly, I prepared CEB dynein and
61 dynactin for preliminary analysis with negative stain EM (Figure 3-1 A & C). The ultimate goal of this collaboration was to obtain cryo-EM structures of both particles, which would require concentrations of around 1-2 µM (1-2 mg/ml for a 1
MDa complex). Unfortunately, we found the yield of CEB protein to be insufficient for large scale sample preparations trial, primarily due to protein loss during concentration in Millipore spin concentrators. To facilitate progress on this project, we turned to bovine material to identify conditions that would allow preparation of cryo-EM grids, operating under the assumption that these same conditions could be used later for CEB protein.
As a first step, negative stain EM was used to characterize the bovine particles and compare the resulting structures with those from CEB preparations.
This revealed that the structural heterogeneity previously reported for porcine dynactin was not a concern, as the CEB and bovine particles looked identical
(Figure 3-1 A and B). Similarly, there were no notable differences between the bovine and CEB dynein preparations (Figure 3-1 C & D). This allowed us to pursue cryo-EM work using bovine proteins, which can easily be obtained in much higher yields, making it possible to achieve the necessary concentration of
1-2 µM.
In the course of this work, we identified multiple factors that contributed to protein loss and instability. A modification to the published MonoQ purification allowed for an increase in protein yield, without sacrificing purity. Briefly, I used a two-step MonoQ purification in place of the published “salt bump” protocol 3.
62
63 Figure 3-1. Two-dimensional negative stain EM averages of (A) CEB dynactin,
(B) bovine brain dynactin, (C) CEB dynein and (D) bovine brain dynein. Scale bars are 10 µm.
64 This utilized a slow linear salt gradient elution, which allowed for relatively broad peaks of dynein and dynactin to elute (Figure 3-2 A). Although these were not highly concentrated, the peaks were resolved from contaminant proteins that elute between the dynein and dynactin peaks (Figure 3-2 B). After pooling the dynein or dynactin fractions, they were rechromatographed on a MonoQ column using a slow salt gradient into and out of a calculated salt bump (Figure 3-2 C), effectively eluting a concentrated pool of pure dynein or dynactin (Figure 3-2 D &
E).
Even with increased protein yields we observed significant protein loss in the Millipore spin concentrators when following the manufacturers instructions.
We found that the addition of the reducing agent tris (2-carboxyethyl) phosphine
(TCEP), which is more resistant to oxidation than other commonly used reducing agents, to a final concentration of 1 mM, in addition to several brief centrifugation steps, significantly reduced protein loss. Specifically, rather than using the manufacturer’s recommended 20 minute centrifugation, we reduced this to 2 minutes, mixing the resulting concentrated protein with unconcentrated sample.
These short spins were repeated until the desired concentration was obtained.
With regard to protein stability, made the serendipitous observation that dynein is temperature sensitive, appearing to unfold when exposed to >26°C for an extended period of time. The bovine dynein/dynactin purification includes a
17-hour centrifugation run into a sucrose density gradient. During one purification run, an equipment malfunction resulted in the ultracentrifuge not
65
66 Figure 3-2. Modified bovine brain dynein/dynactin purification protocol. (A) The first MonoQ run utilizes a 150 ml (0-500 mM KCl) gradient to resolve dynein and dynactin from contaminant proteins, as seen in Coomassie Blue stained SDS-
PAGE analysis (B). (C) Measuring the conductivity at which the dynein and dynactin complexes begin to elute. This conductivity measurement is compared with buffer containing 0 M and 1 M KCl to determine the required KCl concentration. A slow gradient is then used to reach the determined KCl bump, allowing for contaminant proteins to elute separately from the dynein (D) and dynactin (E), which elute as a concentrated peak.
67 holding a vacuum, causing a temperature increase. Unlike samples maintained at 4°C, dynein particles that experienced high temperatures appeared to unfold into elongated structures. Only the heavy chain sediments at ≈20 S in these conditions, as observed with Coomassie blue stained SDS-PAGE (Figure 3-3 A
& B). Presumably, the other dynein subunits sedimented as smaller particles in the sucrose gradient, as was shown for the IC-LC complex (≈5 S) 59. It is likely that smaller particles were not observed because only fractions corresponding to
> ≈17 S were analyzed. The ≈20 S particles appeared as long worm-like structures when examined with negative stain EM, (Figure 3-3 C), suggesting that the HC had unfolded. Dynactin recovered from the same sucrose gradients showed no notable differences, indicating that it is considerably more stable that dynein.
EM analysis of the dynactin complex:
We initiated cryo-EM trials using dynactin, as it shows considerably less conformational heterogeneity than dynein. Particle uniformity facilitates averaging and structural determination, as there are fewer conformations to account for. Like the grids used for negative stain EM, cryo-EM grids are carbon coated. This coating can be charged to allow for better distribution of the sample across the surface of the grid. However, the carbon can also absorb or scatter the electron beam, interfering with high-resolution analysis. To circumvent this problem, cryo-EM grids are designed with holes in the carbon film (Figure 3-4 A).
To be suitable for high resolution structure analysis, the sample must be a single
68
69
Figure 3-3. Analysis of dynein exposed to elevated temperatures. Coomassie
Blue stained SDS-PAGE analysis of sucrose density gradient fractions that were sedimented at 4° C (A) and 26° C (B). The red and green boxes in (B) indicate the expected position of the IC and LIC proteins, respectively. (C) Analysis of the resulting ≈20 S sample shows an extended particle that does not resemble the native HC.
70 layer of protein that is evenly dispersed within an aqueous film within the holes.
This aqueous film is rapidly frozen then analyzed. Ideally the particles will be oriented in all possible orientations to allow a complete three-dimensional reconstruction. At concentrations < 1 µM, the dynactin aggregated at the edge of the holes, due to preferential interactions with the carbon-coated portions of the grid. At concentrations > 1 µM, the protein was distributed throughout the aqueous film, but most of the particles were still aggregated at the edges (Figure
3-4 B). To circumvent this problem, amphiphilic polymers (amphipols) were used to optimize the dispersion of the dynactin particles. The amphipol structure, in which hydrophobic chains are linked to a strongly hydrophilic backbone, allows them to be used in place of detergents to solubilize membrane proteins (Figure
3-4 D) 168. They are also useful for cryo-EM work, where detergent is problematic, as micelles cause ice of variable thickness to form 169. The use of amphipols in EM is relatively new and has had profound impact on the types of molecules that can be imaged in the frozen hydrated state. Amphipol A8-35
(0.0025% w/v) caused the dynactin particles to become evenly spaced in the aqueous film (Figure 3-4 C), and allowed us to obtain data on the large number of particles necessary for image averaging.
Our first observations revealed that most particles lacked the shoulder/arm domain, indicating that it had dissociated from the Arp filament. We reasoned that the complex might be destabilized owing to shipping and/or storage for > 72 hours, but unfortunately this was unavoidable because the samples had to be shipped cross country to The Scripps Research Institute (La Jolla, CA). To
71
72 Figure 3-4. Cryo-EM materials. A) Schematic of EM grids used for Cryo-EM studies. The carbon coated grids have 2 µm holes spaced 2µm apart. Cryo-EM field of dynactin without (B) and with amphipols (C), shows how the presence of the amphipol allows for better particle distribution. (D) Amphipol A8-35 structure, the side chains (X, Y and Z) are covalently linked in random order, with the stoichiometry of X=17, Y=28 and Z=25 168,169. Scale bars are 200 nm.
73 obviate this, cryo-EM grids were prepared as soon as possible after the sample was received, but dissociation of the shoulder/arm from the Arp filament could not be prevented. It has since become clear that the shoulder/arm attachment is salt-sensitive, explaining its dissociation in samples containing ≈350 mM KCl.
Although this prevented us from obtaining a cryo-EM structure of the entire dynactin holo-complex, analysis of the resulting 6.5 Å resolution structure of the Arp filament yielded several novel insights. First, we were able to conclude that Arp filament contains 10 Arp protomers (Figure 3-5 A).
Biochemical data suggests this corresponds with eight Arp1 protomers, one conventional actin and one Arp11. Unfortunately, the resolution of our structure prevented us from being able to determine the location of the conventional actin protomer. We also obtained new information regarding the interactions of the
CapZ α/β heterodimer with the plus-end (“barbed”) of the Arp filament. The C- terminal ≈30 amino acids, termed the “α- and β-tentacles”, of the CapZ heterodimer are positioned identically to what is seen for conventional actin
(Figure 3-5 B & C) 170 171. We were able to identify the pointed end complex subunits, p62, p25 and p27, despite the lower resolution of this part of the structure (presumably due to the intrinsic flexibility of this domain). Finally, we were able to fit the p27 left-handed β-helical heterodimer crystal structure into the pointed end complex density (Figure 3-5 D) 172.
Fitting the cryo-EM structure of the Arp filament into the 24 Å resolution negative stain EM dynactin structure showed clearly that the CapZ dimer is exposed (Figure 3-6 A), contradicting a previous model in which the shoulder
74
75 Figure 3-5. Cryo-EM Arp filament structure. (A) Cryo-EM density map of the Arp filament, fitted with crystal structures of the CapZ α/β and p25/p27 heterodimer and homology models of the Arp proteins. Analysis indicates that there are 10
Arp protomers in the filament (numbered), the barbed end of this filament (B) binds the CapZ heterodimer identically to what is observed in an actin filament
(C) 171. Arrowheads indicate the locations of the α- and β-tentacles of CapZ. (D)
Docking of the p25/p27 heterodimer into the pointed end shows where it makes contacts with Arp11 and p62. The cryo-EM structure was generated from
133,558 particles.
76 domain encapsulates it 122. Subtraction of the Arp filament structure from the negative stain EM density allowed us to develop a better model of the shoulder domain (Figure 3-6 B). The most obvious feature was a quasi-symmetric structure of two bent tube-like structures stacked on top of one another (Figure 3-
6 B). An additional mass, which sat atop the structure was also visible (Figure
3-6 B). Although no detailed structural insight could be gained at this resolution, the EM envelope provided the first indication that the shoulder/arm components, dynamitin, p150Glued and p24, assemble into two distinct structural domains positioned on the Arp filament with apparent two-fold symmetry. The structure is consistent with biochemical work, which showed that dynactin disruption with the chaotropic salt, potassium iodide, yields both a shoulder and shoulder/arm complexes. The former comprises dynamitin and p24 (2:1 stoichiometry), and the latter consists of p150Glued, dynamitin and p24 (2:2:1 stoichiometry) 87.
EM analysis of the dynein complex:
Although we did not pursue a cryo-EM analysis of dynein, our negative stain data provided a wealth of new information regarding dynein structure. As seen in all previous negative stain images, the AAA+ ring domain is easily observed and dominates the images visually (Figure 3-7 A). Class averages generated from fields of particles reveal a range of head spacing and orientations
(Figure 3-7 B). The V-shaped tail domain is difficult to discern in single particle images (Figure 3-7 A), but can be observed in these averages (Figure 3-7 B).
77
78 Figure 3-6. Reconstruction of the dynactin complex structure. (A) Fitting the
Cryo-EM structure of the Arp filament into the 24 Å negative stain model indicates that the Arp filament accounts for the entirety of the 34 nm core of dynactin. The remaining mass is attributed to the shoulder domain, which binds to the side of barbed end of the filament, is a pseudo-symmetric structure (B).
Red and blue pseudo-coloring highlight the two tube-like structures that appear to be structurally symmetric, whereas the green pseudo-colored region appears to be an additional structure that breaks this symmetry. (C) Cartoon representation of the two symmetric tube-like structures. The negative stains structure was generated from 46,734 particles.
79
Because the motor domains dominated the averages, a different analysis method was required to obtain information of the structure of the more flexible regions of dynein. For this, a “mask” is used to obscure the AAA+ domain in class averages. Regions directly adjacent to the “mask” can be selected then averaged, using the same automated particle averaging procedure used to generate class averages of the entire particle (Figure 3-7 C). This yields a new set of averaged structures, in which less predominant domains become visible.
This process can be repeated multiple times to obtain structural information on regions that exhibit conformational heterogeneity in holo-particles. This so-called focused classification protocol was performed iteratively on the dynein complex, yielding a head structure that was consistent with previous reports, plus important new structural information on the tail and linker domains.
Previous negative stain EM work focused on analysis of the dynein HC linker domain that was “docked” on the AAA+ domains. Our focused classification work allowed for a much longer portion of the HC to be detailed
(Figure 3-8 A). A novel feature that emerged was a nearly 90° bend in the HC that was observed in all averages at a consistent position between the tail domain and the AAA+ domain (Figure 3-8 A, yellow arrowhead). This structural feature may act similarly to a shock absorber that allows dramatic conformational changes of the linker relative to the AAA+ ring to occur, without causing significant movement of the tail domain. Analysis of the distance between the
AAA+ domain and this bend categorizes dynein into two groups. In the first, this distance measures ≈10 nm, and a density can be observed crossing the face of
80
81
Figure 3-7. Focused classification processing procedure. Single particle (A) and averaged (100,000 particles; B) negative stain EM analysis of dynein show the
AAA+ ring domain as the prominent feature (A - yellow arrowheads). Masking and focused classification (C) allows for analysis of domains obscured in both single particle and averaged images.
82 the AAA+ ring (Figure 3-8 B). In the second group, the distance is slightly longer,
≈18 nm, and the linker appears to be fully extended and not interacting with the face of the ring domain such that it is “undocked” (Figure 3-8 C). Linker undocking is likely an EM artifact that represents a non-physiological conformation.
Previous EM analysis showed the dynein tail to be V-shaped but did not provide additional detail. Using focused classification, we were able to localize the IC β-propeller domains and the LIC RAS-like domain (Figure 3-8 D, blue and orange arrowheads). These subunits have been shown to bind AA 446-
701 and AA 649-800 on the HC 58, consistent with their apparent locations in the
EM. We also observed two masses that had not been described previously
(Figure 3-8 D, red and white arrowheads), one of which was at the tip of the tail.
Although the identity of this mass could not be determined with negative stain EM, its location matches that of the recently described heavy chain dimerization domain 57. The second mass was proximal to the two dimpled β-propeller domains and in most averages was positioned between the two heavy chains
(Figure 3-8 D, white arrowhead). Although this mass appeared to be unconnected to the rest of the particle, its consistent location strongly suggested it was an authentic dynein component and not an artifact. Additional focused classification yielded two additional masses associated with this structure (Figure
3-8 E). High resolution structures of LC7, LC8 and Tctex could be fitted into
83 84 Figure 3-8. Dynein structure. Focused classified negative stain dynein structure.
Analysis of the linker (A) and tail (D) domain provided structural detail for these domains, showing a near 90° bend in the linker (yellow arrowhead). Some focused classified images of the linker domain show it docked on the AAA+ domain, cartooned in (B), or undocked (C). AAA1-6 are marked in C and D to indicate their orientation respective to the linker. Analysis of the tail (D) shows the localization of the LICs (orange arrowhead), IC β-propeller (blue arrowhead), the HC dimerisation domain (red arrowhead) and an addition mass (whitel arrowhead), which was confirmed to be LC8 with further focused classification
(E) and fitting of LC crystal strucutres. Combining this information yields the structural orginaztion of the dynein motor (F).
85 these densities indicating the three masses are the dynein light chains. Together these results revealed that the N-terminus of the IC chain projects away from the tail (Figure 3-8 F), an unexpected finding. This orientation and the apparent flexibility of the joint between the IC N-terminus and β-propeller domain have profound implications for how dynein and dynactin interact, as I will discuss later in this chapter.
Generation of Dynein-Dynactin-Bicaudal D2 complexes:
Our ultimate goal was to determine the structure of the dynein-dynactin super-complex. The recent discovery that the dynein-dynactin super-complex can be stabilized by scaffolding proteins allowed us to pursue this vigorously
96,97,125. BicD2 AA 25-400 was previously shown to stabilize the dynein-dynactin super-complex, producing a complex with an S-value greater than 20S, as analyzed by sucrose density gradient sedimentation 125. In addition to biochemically stabilizing the dynein-dynactin super-complex, BicD2 AA 25-400 also causes the motor to be super-processive 96,97. Therefore we used this fragment, tagged at its N-terminus with a superfolder-GFP and a Strep-tag
(generously provided by Richard McKenney), which allowed for easy purification and detection via immunoblotting 96.
We first attempted to isolate dynein-dynactin-BicD2 (DDB) super- complexes from purified bovine dynein and dynactin supplemented with recombinant BicD2 fragment using sucrose gradient sedimentation.
Unfortunately, this method was unsuccessful, because although the BicD2
86 fragment shifted into the fractions containing dynein and dynactin, a >20 S complex was not detected (Figure 3-9 A). Furthermore, because sucrose is incompatible with preparation of frozen hydrated samples, we next used size exclusion chromatography (SEC) to isolate DDB super-complexes.
Previous attempts to analyze dynein or dynactin with SEC were plagued by significant protein loss owing to adsorption to the column matrix. For this work, we used very high protein concentrations (0.4 mg/ml dynein, 0.5 mg/ml dynactin; with BicD2 at a ≥ 10x molar excess) and a salt concentration of 150 mM. We were able to recover a larger Stokes radius super-complex (Figure 3-9 B), but we recovered <20 µg/ml in 500 µl, roughly a 5% yield. Assuming a 5% yield, ≈12 mg of dynein and ≈18 mg of dynactin would be required to generate the ≈2.9 mg/ml
(≈1 µM) DDB sample that would be necessary for cryo-EM analysis. Given this unrealistic requirement we abandoned the approach.
Parallel work by our collaborators at the Scripps Research Institute used a different strategy to isolate DDB complexes. Based on the knowledge that cytoplasmic dynein and dynactin bind microtubules in the presence of the non- hydrolyzable ATP analog, AMP-PNP 133 12, they generated native DDB complexes bound to native microtubules. Starting with mouse brains, a high speed supernatant was generated and supplemented with purified tagged BicD2
AA 25-400 (500 nM). Then GTP, taxol and AMP-PNP were added to induce microtubule polymerization and tight binding of the DDB complex. GTP was included to reduced binding of kinesin family motors, thus minimizing other microtubule binding components that might complicate image analysis (Figure 3-
87
88 Figure 3-9. Generating dynein-dynactin-BicD2 complexes in solution. (A) SDS-
PAGE analysis of Sucrose density gradient sedimentation indicate that dynein and dynactin both sediment at ≈20 S (red box), whereas BicD2 sediments in the light sucrose fractions. Combining dynein, dynactin and BicD2 results in BicD2 shifting into the ≈20 S fractions, however, a larger complex is not apparent. (B)
Separating DDB complexes from the individual components can be achieved with SEC (red trace). The DDB complex elutes in the column void (peak 1), whereas dynein (peak 2), dynactin (peak 3) and BicD2 (peak 4) elute in the included column volume (red and blue traces) as determined by SDS-PAGE.
89 10 A). The microtubules and associated proteins were pelleted and the resuspended pellet was analyzed with negative stain EM. The resulting images represented the first images of the DDB complex bound to microtubules. Particle classification and image averaging yielded a well-defined DDB complex structure
(Figure 3-10 B), into which we could fit the previously determined dynein and dynactin structures (Figure 3-10 C). This revealed several key findings. Although the angle between the microtubule and the DDB complex varies, it is always an acute angle, consistent with the direction of movement. The ≈90° bend in the HC is positioned proximal to the microtubule, whereas the dynactin shoulder domain is oriented away from the microtubule. In some averaged images it is possible to observe a short density, likely p150Glued, projecting from the shoulder and wrapping around the dynein tail (Figure 3-10 B, red arrowhead). Unfortunately, this mass cannot be traced to make contacts with either the IC N-terminus or the microtubule, leaving the structure and position of the known p150Glued-IC interaction site in the supercomplex unknown.
Discussion:
Comparative analysis of concurrently generated EM structures:
Contemporaneously with our EM work, Andrew Carter’s lab, at the MRC
Laboratory of Molecular Biology (Cambridge, UK), was working towards the same goal of obtaining high-resolution structures of dynein, dynactin and the dynein-dynactin supercomplex. Despite some differences in source, composition
90
91 Figure 3-10. Generating microtubule bound DDB complexes. Microtubule affinity enriches for the DDB complex, as judged by Coomassie Blue SDS-PAGE analysis (C). EM averaging of DDB particles yields homogenous structures that can be fitted with the previously generated dynein and dynactin EM structures
(D). in some orientations it is possible to observe a mass, likely p150Glued, wrapping around dynactin to the dynein tail (red arrowhead). Combining all structural data generates a model for DDB complexes bound to microtubules (E).
The negative stain structure was generated from 30,758 particles.
92 and preparation methods, similar conclusions were reached. A comparative analysis of the dynactin, dynein and the super-complex structures follows.
Dynactin comparison:
As in our dynactin analysis, the Carter lab generated a cryo-EM structure of dynactin, although they used glutaraldehyde crosslinking to stabilize the complex and ensure that the shoulder/arm was retained 57. The resulting 4.0 Å resolution Arp filament and pointed end complex structure they derived using fitting of known and modeled components is identical to ours. The high resolution of their structure allowed them to localize the convention actin protomer to the pointed-end of the Arp filament (Figure 3-11 A) 57. They also observed loss of the shoulder domain from the Arp filament during sample freezing, but reduction of the salt concentration and glutaraldehyde crosslinking alleviated this. They were able to obtain a 6.3 Å resolution shoulder structure attached to the Arp filament (personal communication).
Comparison of the pointed-end complex structure organization shows the p25/p27 heterodimer fitted the same mass as we observed (Figure 3-11 B). This makes direct contact with the Arp11 subunit, consistent with our crosslinking data
163. An additional mass is present between the p25/p27 heterodimer and Arp filament. This mass is most likely p62 based on previous immunogold labeling with a p62 antibody and EM analysis 112. Although the resolution of this mass in both structures is too low to permit secondary structure assignments, there are a few notable features in both models. The first of these is a previously
93
94 Figure 3-11. Cryo-EM dynactin structure. Cryo-EM analysis of dynactin resulted in a 4.0 Å Arp filament and 6.3 Å shoulder structure (A) 57. Fitting of actin related protein homology models show the position of the conventional actin protomer
(purple) as the penultimate protomer in the Arp filament. The p25/p27 heterodimer (B) is fitted into the pointed end complex, and makes contacts with both Arp11 and the p62 mass. An α-helical domain projects from the core of the p62 mass across Arp11 and contacts actin (arrowhead). The p62 mass and contacts made by the α-helical projection are also observed in our dynactin structure (C). Analysis of the shoulder domain indicates that is composed of three-helix bundles (D). These bundles appear to be similar in structure (E), although they are not perfectly symmetric. An additional mass abuts these bundles (D and F – green mass), and is composed of six β-strands and two α- helices, and appears to have two-fold symmetry. Although not present in all particles, the projecting arm was docked along the Arp filament (G). These structures show that the projecting arm comprises the N-terminal ≈1050 amino acids of p150Glued (H), and includes two microtubule binding domains, CC1A
(purple), CC1B (yellow) and CC2 (teal) as well as a previously unidentified globular domain, termed the intercoil domain (blue).
95 undescribed α-helix that spans the Arp11 protomer and makes contacts with actin (Figure 3-11 B, arrowhead). Additionally, the p62 mass appears to act as a clamp, wrapping around the p25/p27-Arp filament interface. This matches biochemical data indicating that separate domains of p62 are sufficient for interactions with p25/p27 or Arp11, but incorporation and stable attachment with dynactin requires the entirety of p62 (Brett Scipioni, Ph.D. thesis).
The 6.3 Å resolution of the shoulder domain in the Carter lab structure provides additional structural insight, revealing α-helices that form several three- helix bundles (Figure 3-11 C & D) 57. The organization of these matches the bent tube like structures observed in our structure 60, and shows an additional mass, termed the β-saddle domain, that consists of six β-strands and four α-helices
(Figure 3-11 E). Unfortunately, the 6.3 Å resolution prevented determination of what amino acids contribute to the three-helix bundles and β-saddle domain 57.
A remarkable finding in the Carter lab structures was that a subset of averaged particles included the dynactin projecting arm (Figure 3-11 F) 57. This unambiguously demonstrates that p150Glued AA ≈1-1050 represents the projecting arm, leaving the remaining C-terminal ≈200 amino acids to contribute to the shoulder. This contradicts previous models where the projecting arm was believed to be only AA ≈1-550 of p150Glued 49,87,112,122,160. Furthermore, this identifies a previously undescribed, solvent exposed, globular domain, termed the “intercoil domain” (ICD). This comprises AA 550-930 and is likely the mass previously attributed to the N-terminal domain in platinum coated, freeze etched
EM samples 112. Finally these averages show that AA ≈220-550 (Coiled-coil 1)
96 contains a hinge region, which allows the coiled-coil domain to fold back on itself.
This hinge likely maps to AA ≈350-400 where the predicted coiled-coil propensity is low.
Dynein tail comparison:
To obviate issues arising from dynein’s flexibility, Andrew Carter’s group utilized a recombinant HC fragment (AA 1-1074) which contained binding sites for the IC and LIC subunits, to generate recombinant dynein tail for use in cryo-
EM 57. IC, LIC and LCs binding was confirmed by SDS-PAGE prior to EM analysis, but only the mass corresponding with the IC could be identified (Figure
3-12 A, blue pseudo-coloring). Likely, the LCs were not resolved due to the flexibility of the IC N-terminus, to which they bind. It is less clear why the LIC could not be observed (Figure 3-8 E), but may potentially arise from increased conformational heterogeneity in this region, as there are no additional protein- protein interactions to stabilize this domain in one conformation.
As in our structure, a globular mass at the tip of the tail was easily observed and appeared to mediate HC dimerization (Figure 3-12 A, red pseudo- coloring). Crystallization of a smaller HC N-terminal fragment, AA 1-557, allowed the Carter lab to solve the structure of this dimerization domain to 5 Å. This revealed that AA 1-178 forms a globular structure of mixed α-helices and β- strands (Figure 3-12 C). The remainder of this fragment, AA 181-410, forms an eight α-helix bundle each of which sits beside the dimerization domain (Figure 3-
12 D & E).
97
98 Figure 3-12. Structure of the dynein tail. (A) Cryo-EM analysis of the recombinant tail complex shows the positions of the IC β-propeller (blue) and HC dimerization (red) domains. Crystallographic analysis of the extreme N-terminus of the HC show that the dimerization domain is mixed α-helical and β-strand structure (B). The core of this structure shows two-fold symmetry and contains
β-strands encapsulated by α-helices. On either side of the core are symmetric α- helical bundles (D), which appear to have a hinge region that may provide flexibility in this domain.
99 Dynein-dynactin-Bicaudal D2 complex analysis
The Carter lab used this same dynein tail construct with porcine dynactin and BicD2 (AA 1-400) to generate a dynein tail-dynactin-BicD2 (TDB) complex.
Despite the difference in sample preparation, the resulting structure of the dynein tail complexed with dynactin and BicD2 is markedly similar to ours, showing
BicD2 binding along the length of the Arp filament and the dynein tail domain
(Figure 3-10 C & D and Figure 3-13 A-C). The TDB structure reveals an asymmetry imparted on the HCs (Figure 3-13 A, arrowhead), which I propose is translated through the linker to properly space the dynein motor domains to prevent the autoinhibited state (discussed in Chapter One) and allow for processive motility.
Analysis of a TDB structure generated with a N-terminally tagged BicD2 fragment shows the orientation of BicD2 in the super-complex, indicating that the
BicD2 C-terminus extends beyond the pointed-end complex (Figure 3-13 B & C).
This insight, combined with our microtubule bound DDB structure (Figure 3-10 B
& C), indicates that the cargo-binding domain of BicD2 is positioned distal to the microtubule in the microtubule bound complex.
Structural implications into the regulatory mechanisms of the dynein motor
The DDB and TDB complexes provide the first structures of the dynein- dynactin super-complex, but important questions remain unanswered. Notably, the p150Glued projecting arm and IC N-terminus are not observed in the stabilized complex. Some views of our DDB complex show a small protrusion from the
100
101 Figure 3-13. Structure of the dynein tail-dynactin-BicD2 complex. (A) Cryo EM analysis of the TDB complex shows that binding of the dynein tail domain as part of the larger complex imparts an asymmetry originating in the dimerization domain (arrowhead). (B) Difference mapping of TDB complexes generated with a
GFP-tagged and untagged BicD2 fragment determine the orientation of BicD2.
This mapping indicates that the BicD2 N-terminus binds near the barbed end of dynactin, while the C-terminus projects away from the pointed end, allowing it to bind cargo (C).
102 shoulder domain, likely the p150Glued projecting arm, but this does not contact the
IC or microtubule (Figure 3-10 B, arrowhead). Given that we are unable to detect this interaction once the supercomplex has formed and is docked on the microtubule, despite functional assays demonstrating the necessity of this interaction for dynein-based motility 113,114,117,165,167, we believe it plays a role only in the initial stages of DDB complex formation.
Although the TDB complex demonstrates that the super-complex can be formed in solution in the absence of microtubules, it is unclear whether targeting dynein and dynactin to microtubules facilitates assembly in vivo (Figure 3-14 A).
We propose a model in which dynactin is targeted to the microtubule via microtubule end-binding proteins, such as EB1 and CLIP170. In the microtubule bound state an interaction between p150Glued and the dynein IC forms a loosely associated dynein-dynactin complex (Figure 3-14 B), potentially allowing for non- processive movements. When this loosely associated dynein-dynactin complex is in proximity to BicD2, or other scaffolds, interactions among the scaffold, dynein and dynactin would lock the dynein-dynactin-scaffold (DDS) complex in an active state, producing ultra-processive motility (Figure 3-14 C). The DDS complex would be able to bind cargo through either or both dynactin or the scaffold protein.
The insight provided by these structural studies raises several additional questions. We assume that different scaffolding proteins, such as Fip3, Spindly or the Hook proteins, will produce dynein-dynactin complexes that have similar structures to the DDB complex, because the confirmed scaffold proteins contain
103
104 Figure 3-14. Loading model of the dynein-dynactin-scaffold protein supercomplex. (A) Dynein and dynactin are able to bind microtubules, but do no undergo processive transport. (B) A primed dynein-dynactin motor is generated via an interaction between the p150Glued arm of dynactin and the N-terminus of the dynein IC, this primed motor is able to undergo some motility, but is not highly-processive. (C) The primed dynein-dynactin motor is transformed when it interacts with a scaffold protein, locking the dynein-dynactin complex into an ultra-processive motor, which is able to bind and transport cargo.
105 a coiled-coil domain that is predicted to be responsible for interacting with dynein and dynactin. However, this needs to be shown definitively. Additionally, the various scaffold proteins interact with different subsets of organelles, presumably providing for cargo selection specificity and regulation of the dynein-dynactin motor. In addition to lacking structural information on the various DDS complexes, the specific protein-protein interaction that provide for the stepwise assembly remain unknown. Furthermore, whether interactions with different scaffolds occur by chance, with the scaffold and cargo selected due to their proximity to a loosely associated dynein-dynactin complex bound to a microtubule, or if scaffold selection is a regulated process, needs to be determined. Combining structural work, in vitro motility assays with purified components and live cell imaging assays will answer these questions.
Materials and Methods:
Chick embryo brain dynein and dynactin purifications:
Chick embryo brain dynein and dynactin were purified with slight modifications from the published protocol 12. Briefly, 10-12 day chick embryo brains were harvested and flash frozen in 10 g aliquots. Each protein purification utilized 20 g of brains, these were rapidly thawed at 37°C, then dounce homogenized in homogenation buffer with protease inhibitors. A S100 supernatant was generated, rather than pelleting endogenous microtubules as in the published protocol, polymerized purified bovine PC tubulin was added to a final concentration of 0.5 mg/ml. The S100 supernatant supplemented with
106 polymerized PC tubulin was incubated with 20 µM Paclitaxel, 4 mM AMP-PNP, 4 mM MgSO4 and 1 mM GTP, at 37°C for 30 minutes. This sample was sedimented through a 12.5%/25% sucrose cushion supplemented with 20 µM
Paclitaxel and 1 mM GTP. The resulting microtubule pellet was resuspended in
ATP release buffer, then resedimented. The ATP release supernatant was loaded onto a 5-20% linear sucrose gradient as published 12.
The next day, the 20 S dynein and dynactin containing fractions from the sucrose gradient were determined by Coomassie blue stained SDS-PAGE.
These fractions were pooled and diluted 2 fold with TM buffer. This diluted sample was loaded onto a MonoQ 5/50 GL column (GE Life Sciences), washed until the baseline returned to zero. The protein was eluted with the following gradient:
15 ml linear gradient from 0-200 mM KCl
Step from 200-250 mM KCl, remain at 250 mM KCl for 2.5 ml
8 ml linear gradient from 250-280 mM KCl
Step from 280-310 mM KCl
25 ml linear gradient from 310-500 mM KCl
10 ml linear gradient from 0.5-1 M KCl.
Fractions containing dynein and dynactin were assessed for purity with
Coomassie Blue Stained SDS-PAGE, then pooled and supplemented with 1 mM
ATP and 1 mM DTT. If necessary, the dynein and dynactin were concentrated using an Amicon Ultra 0.5 ml Centrifugal Filter (EMD Millipore) with a 30 KDa molecular weight cut off.
107 Bovine brain dynein and dynactin purification:
Bovine brain dynein and dynactin were purified as published with modifications to the MonoQ elution profile 3. To ensure a higher purity of the final bovine dynein and dynactin samples, the final MonoQ chromatography purification step was altered such that the 20 S dynein and dynactin containing fractions from the sucrose gradients were loaded onto a MonoQ HR 5/5 column
(GE Life Sciences). This protein was eluted using a linear salt gradient from 0-
1M KCl in TM buffer, 35 mM Tris, 5 mM MgSO4, pH 7.2, over 50 ml, collecting 1 ml fractions across the elution gradient. The fractions containing dynein and dynactin were pooled separately, and diluted to four time the volume to reduce the salt concentration. The conductivity of the first and last fraction pooled was used to determine the concentration of salt at which the protein eluted.
The dynactin pool was then loaded onto a 1 ml MonoQ GL 5/50 column
(GE Life Sciences), washed until the baseline returned to zero. A 15 ml linear gradient was used to reach the calculated KCl concentration for the salt bump, generally around 320 mM KCl to 400 mM KCl. The flow rate was reduced to
0.25 ml/min, and 0.3 ml fractions were collected for 2 ml, then a 10 ml linear gradient to 1 M KCl was run at 1 ml/min. The column was equilibrated into TM buffer, then the dynein sample was loaded onto the column, and washed until the baseline returned to zero. A 10 ml linear gradient was used to reach the calculated KCl concentration for the dynein salt bump, generally around 225 mM
KCl to 275 mM KCl. The flow rate was reduced to 0.25 ml/min, and 0.3 ml fractions were collected for 2 ml, then a 15 ml linear gradient to 1 M KCl was run
108 at 1 ml/min. Peak fractions for both dynein and dynactin were supplemented with
1 mM ATP and 1 mM DTT and assessed with Coomassie Blue stained SDS-
PAGE, then pooled and concentrated further in an Amicon Ultra 0.5 ml
Centrifugal Filter (EMD Millipore) with a 30 KDa molecular weight cut off if needed.
Bicaudal D2 fragment purification:
The BicD2 AA 25-400 fragment, which had an N-terminal His6X-StrepII-
Superfold GFP tag,(from R.J. McKenney – R.D. Vale lab) was grown to an OD600 of 0.4-0.6 at 37°C, induced with 1 mM IPTG and grown overnight at 18°C. After centrifugation, the bacterial pellet was resuspended in 50 mM Sodium Phosphate,
300 mM NaCl, 1 mM TCEP with protease inhibitors, pH 8.0. The resuspended pellet was lysed with an Avestin EmulsiFlex C3 then centrifuged to remove the insoluble debris. The supernatant was filtered through a 0.2 µm syringe filter and combined with 5 ml of Strep-Tactin Superflow Plus resin (Qiagen) and incubated for 2 hours at 4°C. The resin was pelleted using a dynac table top centrifuge, then washed with 50 mM Sodium phosphate, 300 mM NaCl, 1 mM TCEP with protease inhibitors, pH 8.0. This wash step was repeated 4 additional times; the resin was resuspended in 5 ml elution buffer, 10 mM Desthiobiotin, 50 mM
Sodium Phosphate, 300 mM NaCl, 1 mM TCEP with protease inhibitors, pH 8.0 and incubated for 15 min on ice. The resin was pelleted at 2 K RCF in a
Eppendorf Centrifuge 5417 C. The elution step was repeated a total of four times to allow for maximum protein recovery.
109 After elution, the eluted BicD2 protein was dialyzed into 35 mM Tris, 5 mM
MgSO4, 50 mM KCl, pH 7.2, until buffer exchange was complete. One quarter of the dialyzed protein was loaded onto a MonoQ 5/50 GL (GE Life Sciences), the remaining protein was snap frozen in 5 ml aliquots for later use. The column was washed until the baseline returned to zero, then eluted using a 50 ml linear gradient from 50 mM KCl to 1 M KCl in 35 mM Tris, 5 mM MgSO4, pH 7.2, with 1 ml fractions collected. Peak fractions were analyzed for purity with Coomassie
Blue Stained SDS-PAGE, then pooled and snap frozen in 0.75 ml aliquots.
Immediately prior to use, one 0.75 ml BicD2 aliquot was thawed, filtered with a 0.2 µm syringe filter and 0.5 ml was loaded onto a pre-equilibrated
Superose12 10/300 GL column (GE Life Sciences) in 35 mM Tris, 5 mM MgSO4,
150 mM KCl. The column was run at 0.25 ml/min, with 0.5 ml fractions collected.
BicD2 AA 25-400 eluted separately from contaminating proteins, within the included volume of the sizing column.
DDB complex assembly:
Dynein-dynactin-BicD2 AA 25-400 complexes assembly was initially analyzed using SEC. To generate complexes, 150 µg of dynein was combined with 225 µg of dynactin, with or without 60 µg of BicD2 AA 25-400 and brought to a final volume of 0.65 ml. As the purified dynein and dynactin contained relatively high KCl concentrations, ~250 mM and ~375 mM KCl respectively, the dynein-dynactin or dynein-dynactin-BicD2 solutions were dialyzed against 35 mM
Tris, 5 mM MgSO4, 150 mM KCl for 1 hour at 4°C. The dialyzed protein sample
110 was filtered through a 2 µm syringe filter, then loaded onto the pre-equilibrated
Superose12 10/300 GL column (GE Life Sciences). The column was run at 0.25 ml/min, with 0.5 ml fractions collected during the entirety of the run. Fractions of interest were determined by absorbance (280 nm) measurements, 300 µl of these fractions were TCA precipitated and further analyzed for protein composition with Coomassie Blue Stained SDS-PAGE.
As SEC was not a efficient means of generating DDB complexes, we next turned to a microtubule affinity method to enrich for the DDB complexes 60.
Briefly, a modified CEB protocol was used to enrich for the DDB complexes, such that a S100 mouse brain supernatant was supplemented with 500 nM BicD2 AA
25-400, incubated with gentle agitation for 2 hours on ice. This was further supplemented with 4 mM MgSO4, 4 mM AMP-PNP, 1mM GTP and 20 µM taxol, then incubated for 25 min at 37°C. Microtubules and associated proteins were pelleted and gently resuspended in 35 mM PIPES, 35 mM KOH, 5 mM MgSO4, 1 mM EGTA, 0.5 mM EDTA, pH 7.0 with 1 mM GTP, 4 mM AMP-PNP, 4 mM
60 MgSO4 and 20 µM taxol then subjected to negative stain EM as described .
111
Chapter Four
Mapping of the dynactin shoulder domain aimed at
understanding its three-helix bundle structure
112
Introduction:
Dynactin was initially identified as an activator of the microtubule based motor cytoplasmic dynein (hereafter dynein) 12,84. Further work showed this dynein activator is composed of 11 subunits, with stoichiometric ratios ranging from one to eight subunits per dynactin molecule84,85,87,112,173-176. These subunit stoichiometries result in structural asymmetry within the dynactin complex, which contrasts the compositional and structural symmetry of the homodimeric dynein complex. Although dynactin as a whole exhibits asymmetry, one structural domain, the shoulder/arm complex, is compositionally symmetric, as it contains dynamitin, p24 and p150Glued in a four to two to two ratio87.
Overexpression of full length dynamitin or a N-terminal fragment, amino acids 1-87 (hereafter N87) in vivo releases the entire shoulder/arm complex by replacing the endogenous dynamitin bound along the Arp filament 121,177. These results can be reproduced in vitro with purified components, indicating that dynamitin is the causative agent and does not require other cellular factors. The release caused by excess N87 implicates the N-terminus of dynamitin as the domain responsible for anchoring the shoulder/arm complex to the Arp filament, which is further supported by biochemical work showing that N87 binds specifically and directly to Arp1, but not actin 121.
A recent high-resolution electron microscopy study supports the hypothesis that the N-terminus of dynamitin binds along the Arp filament and showed that the remainder of the shoulder domain is largely α-helical and
113 exhibits mirror symmetry 57. This structure contains two U-shaped three-helix bundles, where one arm of the U is about twice the length of the other arm.
These two structures stack on top of each other with the long arms on opposite sides. In the center of this structure is a mixed α-helical/β-strand structure, termed the β-saddle domain 57. Two additional densities are seen projecting away from the end of each long arm of the U-shaped structures. Although the density is not continuous with the U-shaped structure, these densities are also consistent with three-helix bundles.
The high-resolution EM reconstruction of dynactin allowed for unambiguous identification of all dynactin protomers, with the exception of dynamitin, p24 and AA ≈1050-1280 of p150Glued. Although the resolution of the shoulder domain was insufficient to assign amino acid identities to the α-helical domains, the structure is sure to include dynamitin, p24 and the p150Glued C- terminus as these are the only proteins unaccounted for. Although the amino acid identities are not known, the U-shaped three-helix bundle is assumed to be a trimer of dynamitin and p24. This is consistent with analysis of native dynactin, which contains a trimeric complex with two dynamitin protomers and a single p24 protomer that can be reconstituted from recombinantly expressed dynamitin and p24 87,177,178. Furthermore, the C-terminus of p150Glued passes between the U- shaped three-helix bundles and is presumed to contribute to the saddle 57.
In addition to forming three-helix bundles with dynamitin, p24 appears to be required for p150Glued incorporation into the dynactin complex. Deletion of the yeast p24 homolog (ldb18) results in the loss of the p150Glued homolog (Nip100)
114 from dynactin, while the dynamitin homolog (Jnm1) remains associated with the
Arp filament179. The notion that p24 provides the critical bridge between dynamitin and p150Glued highlights the importance of obtaining detailed structural information on the shoulder complex, which remains the least defined part of the dynactin structure. The dynamitin-p24 complex can be readily reconstituted from recombinant proteins, which facilitates analysis 177,178. However, neither recombinant dynamitin nor p24 behave ideally when expressed in bacteria. Full- length dynamitin and shorter dynamitin fragments are readily soluble, but form large oligomers178. Full-length p24 is insoluble when expressed in bacteria, but can be renatured in the presence of dynamitin to form a soluble, well-behaved hetero-trimeric complex, with a two to one, dynamitin to p24 stoichiometry. In the work described in this chapter, I took advantage of the ability of dynamitin to solubilize p24 and the ability of p24 to suppress dynamitin oligomerization to define minimal interaction domains. This work allows me to propose a structural model of the unusual bent three-helix bundle that comprises much of the mass of the dynactin shoulder 57.
Results:
Dynamitin and p24 sequence analysis:
Sequence analysis and circular dichroism show dynamitin contains both structured and intrinsically disordered regions 121. The N-terminal 87 amino acids, which are responsible for interactions of the entire shoulder/arm complex with the
Arp filament are unstructured in solution but become structured when bound to
115 the Arp filament 57,121. The rest of dynamitin is predicted to form a series of α- helices (PSIPRED, Figure 4-1 B), which are interrupted by a predicted intrinsically disordered “hinge” domain, AA 210-215, (PONDR-FIT, DISOPRED2 and GlobProt2). This boundary divides dynamitin into roughly two halves, with the entire C-terminal portion being predicted to form coiled-coil or multi-coiled high-order oligomers 178 (Multicoil and Paircoils2; Figure 4-1C). The dynamitin N- terminal portion contains a predicted α-helical domain spanning AA 103-190, with
AA 103-139 predicted to be coiled-coil or multi-coil. This domain forms trimeric or tetrameric complexes in vitro 178, but does not oligomerize further. These structural predictions were used to design truncated dynamitin fragments to map the domains that interact with p24 (Figure 4-2 A).
Similar analysis predicts that p24 is highly α-helical (PSIPRED). Although the amino acid sequence of p24 is not well conserved, the boundaries of the predicted α-helices are consistent (Figure 4-1 D-F). Therefore, these boundaries were used to design a series of truncated p24 fragments (Figure 4-2 B) to be used to further map p24-dynamitin interactions. The resulting p24 fragments are predicted to be strongly α-helical, but only two helices are predicted to have coiled-coil or multi-coil propensity (Multicoil and Paircoils2; Figure 4-1 F). The first predicted multi-coil spans amino acids 40-65 and the second is closer to the
C-terminus, spanning amino acids 125-160. Using both full length and fragments of dynamitin and p24 I mapped the interaction domains between these two proteins within dynactin’s shoulder.
116
117 Figure 4-1. Sequence analysis of dynamitin and p24. Conservation of the dynamitin (A) and p24 (D) amino acid sequence. Predicted α-helical domains
(blue highlight) for the dynamitin (B) and p24 (E) amino acid sequences.
Predicted coiled-coil (light green highlight) and multi-coil (dark green highlight) domains for the dynamitin (C) and p24 F) amino acid sequences.
118
119 Figure 4-2. Dynamitin and p24 fragments. (A) Dynamitin (DM) fragments used to map interactions with p24 fragments (B). Summary of solubilization of full-length p24 with DM fragments (top right) and of p24 fragment solubility after renaturation alone or with full length DM (bottom middle).
120 Solubilization of p24 fragments with full length dynamitin:
Previous studies developed an assay to evaluate dynamitin-p24 interactions, based on the ability of full-length dynamitin to restore the solubility of full-length p24, a protein that is ordinarily insoluble 178 177. I used this assay to obtain preliminary information regarding the minimal dynamitin fragment needed to interact with full-length p24. I found that amino acids 33-186 of p24 were soluble on their own, although this fragment formed large oligomers that eluted in the column void of the size exclusion chromatography (SEC) column (Figure 4-3
A and B). p24 fragments containing amino acids 1-32 (i.e. amino acids 1-67 and
1-108) could not be solubilized even when reconstituted with full length dynamitin
(Figure 4-3 C and D), suggesting that AA 1-32 of p24 do not interact with dynamitin.
Because the p24 fragments lacking AA 1-32 could be recovered as soluble proteins in the absence of dynamitin, simply renaturing p24 and dynamitin fragments together does not reflect an interaction. Therefore, all further work included a step in which complexes of dynamitin and p24 were isolated on a His-Trap column utilizing the 6X-His-tag on p24, prior to further characterization with SEC.
Dynamitin’s N-terminus (AA 1-212) is not necessary for complex formation:
Dynamitin’s N-terminus (AA 1-87) binds to Arp1. I verified that this dynamitin domain is not necessary for interactions with p24 by reconstituting
121
122 Figure 4-3. The p24 N-terminal 32 amino acids interfere with complex formation with dynamitin. Coomassie Blue stained SDS-polyacrylamide gel showing the supernatant (S) and pellet (P) for renatured p24 AA 33-186 alone (A), p24 AA 1-
67 with Full length DM (C) and p24 AA 1-108 with full length DM (D). (B) SEC analysis of p24 AA 33-186 and Coomassie Blue stain SDS-polyacrylamide gel of peak fractions.
123 complexes using dynamitin that lacked this portion. As expected, dynamitin AA
88-406 formed a complex with full-length p24 (Figure 4-4 B). Densitometric analysis of SyproRuby stained gels revealed a 2:1 stoichiometry (Figure 4-4 D), like full length dynamitin with full-length p24. The dynamitin-p24 complex had a
Stokes radius of 52.2 ± 2.1 Å, as compared to 62.5 ± 1.5 Å for dynamitin AA 88-
406 alone (Figure 4-4 C). This size shift, which is also seen when full length dynamitin is complexed with full length p24 (66.6 ± 0.9 Å vs. 59.2 ± 0.2 Å; Figure
4-4 A and B), may indicate that the dynamitin-p24 complex undergoes structural compaction.
The cryo-EM structure of dynactin reveals that the shoulder (i.e. dynamitin and p24) is largely composed of three-helix bundles 57. Dynamitin AA 88-406 is
319 amino acids whereas p24 is only 186 amino acids, which is inconsistent with a simple longitudinal assembly in which the proteins are aligned along their lengths. To learn more about which regions of dynamitin and p24 might comprise the α-helical bundle in the shoulder, I used the solubilization assay to survey complex formation between different dynamitin and p24 fragments. This analysis was performed with both full length p24 and p24 AA 33-186. The latter version was included based upon my earlier results where the presence of p24
AA 1-32 was incompatible with complex formation. These p24 fragments were combined with dynamitin fragments truncated from the N-terminus to begin to define the minimal dynamitin fragment sufficient to support interaction p24.
I had previously shown that AA 213-406 did not rescue full-length p24 solubility 178, so I was surprised to find that this fragment, with the N-terminal His-
124
125 Figure 4-4. The N-terminal 212 amino acids expendable for formation of dynamitin-p24 complexes. Chromatogram of SEC analysis of full-length dynamitin (DM) (A), the DM – p24 complex, DM AA 88-406 alone (C), the DM 88-
406 – p24 complex, (D), DM)AA 213-406 alone (E) and the DM 213-406 – p24 complex (E). Stokes’ radius and stoichiometry, as measured by densitometric analysis with SyproRuby, are indicated. Coomassie Blue SDS-polyacrylamide gel showing peak fractions.
126 tag removed, supported complex formation with both full-length p24 and p24 AA
33-186. I observed nearly identical elution behavior for the dynamitin-p24 complex compared with dynamitin alone (52.4 ± 0.4 Å vs. 53.9 ± 0.4 Å; Figure 4-
4 C and D). The length of these fragments is more consistent with a longitudinal assembly model, with the dynamitin fragment containing 194 amino acids and the p24 fragment consisting of 154 amino acids. Furthermore, these data indicate that dynamitin AA 1-212 are not required for p24 binding and are unlikely to contribute to the bent three-helix bundle in the dynactin shoulder. Given this, I propose that AA 88-212 contributes to the masses designated the “paddle” and
“hook” domain in the cryo-EM structure 57 (Figure 4-6, C and D), as these are the only remaining masses that can accommodate this part of dynamitin.
Dynamitin’s C-terminus is necessary for complex formation:
A dynamitin fragment truncated from the C-terminus, AA 88-310, showed no evidence of complex formation with full-length p24, as the dynamitin failed to bind and elute from the HisTrap column with p24 (Figure 4-5 B). To determine if p24 AA 1-32 was interfering with complex formation I also examined p24 AA 33-
186. This showed identical behavior to the full length protein in this set of experiments (Figure 4-5 D), forming complexes with dynamitin AA 88-406 (Figure
4-5 A and C), but not the C-terminal truncated fragment, AA 88-310. This result indicates that the last predicted helix in dynamitin, which has strong multi-coil propensity, is required for complex formation with p24.
127
128 Figure 4-5. The C-terminal 97 amino acids are necessary for dynamitin-p24 complex formation. Chromatogram of the HisTrap purification of renatured dynamitin (DM) 88-406 with full-length p24 (A), DM 88-310 with full-length p24
(B), DM 88-406 with p24 AA 33-186 (C) and DM 88-310 with p24 AA 33-186 (D).
Coomassie Blue SDS-polyacrylamide gel showing the column load (L), flow- through (F) and elution (E) fractions for each purification.
129 This demonstrates that C-terminal 194 amino acids (AA213-406) of dynamitin is the domain that interacts with p24. Although the lengths of these protein fragments are not identical (194 and 154 amino acids respectively) it is possible that the two dynamitin molecules wrap in a tight coiled-coil, with the p24 molecule forming a three-helix bundle with only portions of this structure.
Discussion:
Electron microscopy studies have identified the location of most dynactin subunits 57,112 60. The exception to this is the components of the shoulder domain.
This has a primarily α-helical structure, but amino acid identities of the α-helices could not be determined, preventing rigorous identification of the components 57.
The shoulder domain appears to be four portions: two short three-helix bundle structures, termed the hook and paddle, a longer U-shaped three-helix bundle structure and a mixed α-helical/β-strand domain, termed the saddle (Figure 4-6
A). The three-helix bundle structures are present in duplicate and show mirror symmetry, with the p150Gliued C-terminus passing between them.
I showed that dynamitin AA 213-406 forms a trimeric complex with p24 AA
33-186 that is presumed to be a three-helix bundle (Figure 4-6 E and F). It is not known whether this α-helical bundle is a parallel or anti-parallel structure.
Regardless of the binding orientation, the length of dynamitin-p24 complex can only be accommodated in the U-shaped three-helix bundle, because the other α- helical domains, the hook and paddle 57, are too small to accommodate it.
130
131 Figure 4-6. Structure and polypeptide identity of the shoulder domain. (A)
Structure of the dynactin shoulder, thick bars represent α-helices, thin lines are random coil and arrows represent β-strands. (B) The β-saddle domain of the dynactin shoulder which is presumed to be largely p150Glued with one α-helix contributed from the p24 N-terminus. The paddle (C) and hook (D) domains are nearly identical three α-helical structures believed to be dynamitin AA 88-212.
The bent three-helix bundle structures (E and F) that are identified as p24 AA 33-
186 and dynamitin AA 213-406.
132
Assignment of dynamitin AA 213-406 and p24 AA 33-186 to the bent three-helix bundle allows me to make predictions regarding the identity of the proteins in the remaining shoulder structures. The cryo-EM structure shows the
N-termini of dynamitin bound to the Arp filament, and appearing to terminate near the hook and paddle domains 57. The structural similarity between the hook and paddle, both of which are three-helix bundles, with two short α-helices (≈20 AA each) and a longer α-helix (≈40 AA), suggests that these two domains may be compositionally the same (Figure 4-6 C and D). The remaining portion of dynamitin, AA 89-212, is predicted to be α-helical and to adopt a spectrin-like fold
(SWISS-MODEL). Given that there are four dynamitin protomers present in dynactin, this prediction would be consistent with the observed structure.
The cryo-EM structure also shows the site where the p150Glued arm projects from the shoulder domain. This is a coiled-coil, comprising AA 926-
1048 of p150Glued. Within the shoulder the p150Glued dimer can be seen to separate into two α-helices, consistent with the predicted structure of p150Glued
AA 1048-1094 (Multicoil and Paircoils2), and can be seen leading towards the hook domain, where the α-helical density terminates 57. The remainder of the p150Glued C-terminus is predicted to contain long stretches of random coil, several
α-helices (AA 1098-1131, AA 1153-1169, AA 1189-1220, AA 1237-1246 and AA
1274-1285) and three short β-strands (AA 1172-1178, AA 1250-1257 and AA
1265-1270). I believe that the three β-strands in each p150Glued polypeptide contribute to the saddle domain. This would account for the six β-strands observed in the EM structure. I predict that the four α-helices derive from the N-
133 terminus of p24 (AA 1-33) and the C-terminus of p150Glued (AA 1189-1220). Such a structure would anchor p150Glued solidly in the shoulder domain and is consistent with the necessity of p24 for p150Glued incorporation 179. The remaining regions of random coil in the p150Glued C-terminus may provide structural flexibility. These would not be expected to be observed in the EM structure, but their flexibility would allow the region to be solvent exposed and available for the numerous post-translational modifications that have been reported within this region of p150Glued (PhosphoSite Plus, dbPTM and PHOSIDA).
These predictions account for the majority of the dynamitin, p150Glued and p24 proteins. Remaining unaccounted for regions are predicted to be primarily random coil, which would act to connect the U-shaped domain to the hook, paddle and saddle domains.
In summary, I have used predicted structural information and biochemical binding studies to map the portions of dynamitin, p150Glued and p24 that correspond with the different should domains. My data indicate that p150Glued, dynamitin and p24 interact to form an intricately woven structure 57. Such a structure would explain how p150Glued is tethered to the Arp filament and provides a mechanism for anchoring p150Glued in a way that can withstand tension during dynein-based movement.
134 Materials and Methods
Cloning:
Fragments of p24 were cloned using the full-length pRSET-p24 vector as a template 178. The fragments were PCR amplified with the appropriate primer combination (Table 1), then ligated into the pRSET-B using BglII and EcoRI restriction enzymes.
Fragments of dynamitin were generated from Cys-lite dynamitin 177. The pBAD-His6x-Sumo-TEV (pBHST – gift from C.M. Kaiser lab) plasmid backbone was PCR amplified as three ~2 kb fragments using primers 7-12 (Table 1).
Dynamitin fragments were generated using combinations of primers 13-21 (Table
1). The three pBHST backbone fragments were assembled with the dynamitin inserts using the Gibson Assemble Cloning Kit (NEB – E5510).
Protein purification:
Full length and fragments of p24 were expressed and purified using a modified protocol for full length p24 178. Bacterial cultures for the fragments were grown at 37°C to an OD600 of 0.4-0.6, induced with 1 mM IPTG then grown at
37°C overnight. After centrifugation, the pellet was resuspended in 100 ml resuspension buffer, 100 mM Tris, 5 mM EDTA, 5 mM DTT, pH 7.0, per liter of bacterial culture, then lysed using an Avestin EmulsiFlex C3. After lysis, the insoluble material was separated from the supernatant with centrifugation. The insoluble pellet was washed with 20 ml of wash buffer, 100 mM Tris, 2 M Urea, 5 mM EDTA, 5 mM DTT, 2% Triton X-100, per liter of bacterial culture, then
135 centrifuged to separate the insoluble and soluble material. This wash step was repeated an additional four times, the final pellet was rinsed with the resuspension buffer to remove detergent, then resuspended in 20 ml of 6 M
Guanidine-HCl, 0.5 M NaCl, 20 mM Sodium Phosphate at pH 7.4. This was loaded onto a 5 ml His-Trap (GE Life Sciences), washed until the baseline returned to zero, then eluted with a pH gradient from pH 7.4 to pH 2.1. Protein purity was assessed with Coomassie Blue stained SDS-PAGE.
Dynamitin fragments were purified as soluble proteins. Bacteria carrying the dynamitin plasmids were grown at 37°C to an OD600 of 0.4-0.6, then induced with 0.1% L(+)Arabanose (Sigma- Aldrich – A3256) and grown overnight at 23°C.
After centrifugation, the pellet was resuspended in 50 mM Sodium Phosphate,
300 mM NaCl, pH 7.4 with protease inhibitors, then lysed using an Avestin
EmulsiFlex C3. The supernatant was loaded onto a 5 ml HisTrap column (GE
Life Sciences), washed until the baseline returned to zero, then eluted with 50 mM Sodium Phosphate, 300 mM NaCl, 500 mM Imidazole, pH 7.4.
The peak fractions, containing ~20-50 mg of protein, was combined with
0.5 mg recombinant His-tagged TEV protease to cleave the His-Sumo tag (gift from V.J. Hilser lab), this was dialyzed against 50 mM Sodium Phosphate, 300 mM NaCl, pH 7.4 overnight at 4°C. The cleaved protein was loaded onto a second 5 ml HisTrap column (GE Life Sciences), which bound the cleaved tag and any uncleaved dynamitin. The flow-through sample containing the cleaved dynamitin was collected and dialyzed into 20 mM Tris, pH 7.4 for 2 hours at 4°C.
The dialyzed flow-through was loaded onto a MonoQ GL 5/50 (GE Life Sciences),
136 washed until the baseline returned to zero, then eluted with a salt gradient from
0-500 mM NaCl. . Protein purity was assessed with Coomassie Blue stained
SDS-PAGE.
Refolding Complexes and complex purification:
Dynamitin-p24 complexes were renatured as previously described 178.
Briefly, p24 fragments were dialyzed against 3 M Guanidine-HCl, 0.5 M Arginine,
20 mM Tris, pH 7.4 at 4°C. Both the p24 fragment and dynamitin fragment were then dialyzed into 2 M Guanidine-HCl, 0.5 M Arginine, 20 mM Tris, pH 7.4 at 4°C.
After dialysis, the p24 and dynamitin fragment were combined, with dynamitin in a molar excess, dialyzed further into 1 M Guanidine-HCl, 0.5 M Arginine, 20 mM
Tris, pH 7.4 at 4°C. The sample was then dialyzed into 0.5 M Arginine, 20 mM
Tris, with 1 mM DTT, pH 7.4 at 4°C, then dialyzed against 20 mM Tris, 100 mM
NaCl, 1 mM DTT, pH 7.4 at 4°C. The dynamitin-p24 sample was finally dialyzed against 50 mM Sodium Phosphate, 300 mM NaCl, 50 mM Imidazole, pH 7.4.
After the final dialysis step, the dynamitin-p24 sample was centrifuged at
14 K RPM in an Eppendorf Centrifuge 5417C for 20 min at 4°C to remove insoluble protein. The supernatant was sterile filtered and loaded onto a 1 ml
HisTrap (GE Life Sciences), free dynamitin was unable to bind the column, while dynamitin-p24 complexes bound via the His-tag on p24. The complexes were eluted with 50 mM Sodium Phosphate, 300 mM NaCl, 500 mM Imidazole, pH 7.4.
The eluted dynamitin-p24 complexes could then be analyzed further.
137 Size exclusion chromatography analysis:
Purified dynamitin-p24 complexes were loaded onto a pre-equilibrated
Superdex200 10/300 GL (GE Life Sciences) with 20 mM Tris, 150 mM NaCl, 1 mM β-Mercaptoethanol, pH 7.4. A 0.5 ml protein sample was injected into the column at 0.25 ml/min, the column continued to run at 0.25 ml/min with 0.5 ml fractions collected throughout the entirety of the column run. Stokes’ radii were calculated using a standard curve generated with standards from both the Gel
Filtration HMW and LMW Calibration kits (GE Life Sciences) according to the protocol for the Superdex200 10/300 GL column.
Fractions from the Superdex200 10/300 GL column were analyzed with
Coomassie Blue stained SDS-PAGE. The peak fraction was used to analyze complex stoichiometry. A standard curve of purified dynamitin and p24 was generated using 10 µg, 5 µg, 1.25 µg, 0.625 µg and 0.3125 µg of both purified dynamitin and p24 fragments. These samples were run on the same gel as a serial dilution of the peak fraction from the SEC analysis. Sypro Ruby staining
(BioRad) was used to determine the amount of dynamitin and p24 in the complexes, which could then be converted into a molar ratio.
138
G C
AG
G
ATGGAGACTG GACC CGGTC CTGGC CGTTCAC AGAGCGTC TAGGAAGC TCGCATTGC GGTTTCGCC TCAGGTCTTC CCAGGCGTTTC
CTTTCCCAGCTTCTTC ATGGCGGACCCTAAATAC CAAAGCAAGGTGCACCAG GAACTACATTCTCGGCCTG TCTGGAGAATATGAGATGC GCCTGTCTC ATAAGTGACAAGGCTGCTAT TCCCTGTAGACCTGCAGAAA TGTATCTGCATCTTCTACAG Sequence TTGGAAGTACAGGTTTTCC TAGAATCAGAACGCAGAAGC AACAGATCTATGGCGGCTCT GGGGAATTCTCATTCCTCTG GCGAGATCTAAGGTGGCTGA CTTGAATTCTCACACCAGGG CCCTTATGCGACTCCTGCAT GTATCTTTATAGTCCTGTCG ATGCAGGAGTCGCATAAGGG CCCAGATCTGAGTACATTGA CCCGAATTCTCAGTATTTAA CCGACAGGACTATAAAGATA GCTTCTGCGTTCTGATTTCA AGGAAAACCTGTACTTCCAA AGGAAAACCTGTACTTCCAA AGGAAAACCTGTACTTCCAA AGGAAAACCTGTACTTCCAA AGGAAAACCTGTACTTCCAA GCTTCTGCGTTCTGATTCTA GCTTCTGCGTTCTGATTCTA GCTTCTGCGTTCTGATTCTA
amino acid 1 amino acid 66 amino acid 32 amino acid 71 amino acid 186 amino acid 107 – – – – – – ForwardAA1 PrimerName DM DMforward 88 AA DMForward AA213 DMForward AA260 DMForward AA311 DMReverse AA212 DMReverse 259AA DMReverse 310AA DMReverse 406AA pBHSTfragment 1For pBHSTfragment 2For pBHSTfragment 3For pBHSTfragment 2Rev pBHSTfragment 3Rev PBHST fragmentPBHST 1Rev Forward Forward Forward Reverse Reverse Reverse
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 No.
139 Table 4-1. PCR primers used to generate p24 and dynamitin fragments.
Combinations of primers 1-6 (forward and reverse primers) were used to generate the various p24 fragments. The pBAD-His6x-Sumo-TEV-DM plasmid was generated via Gibson Assembly, where the backbone was amplified in three fragments using primer sets 7/8, 9/10 and 11/12, which was combined with one
DM fragment generated with primer sets from 13-21.
140
Chapter Five
General Conclusions:
141 Since the discovery of the dynein motor complex, many advances have been made regarding its structure and functions (reviewed in180-182). Genetic screens and biochemical analysis identified the individual components of the dynein motor pathway: the dynein complex (HC, IC, LIC and LCs), dynactin
(p150Glued, p62, dynamitin, Arp1, actin, Arp11, CapZ α/β, p27, p25 and p24),
NudE(L) and Lis1 (discussed in Chapter One) 3,5,6,25,29,30,41-46,63-65,67-69,71,73,76-
81,83,87-91,112,132-144. However, the structure of the complex remained less well understood, especially the regulation of how the different components come together to form a functional motor and the precise organization and interactions of subunits within each individual complex. Low-resolution EM analysis provided a general maps of dynein and dynactin 4,112,122,160,161, while additional EM and crystal studies giving more insight into the dynein AAA+ ring domain, microtubule- binding stalk and a small portion of the linker 50-55.
X-ray crystallography studies focused on the dynein motor domain resolved how ATP hydrolysis results in the power-stroke required for motility
(described in Chapter One) 52,183. Additional work demonstrated that the binding of Lis1, with or without NudE(L), generates a stalled, high-force motor that decouples ATP hydrolysis from the conformational changes required to produce the linker swing and yield the power-stroke 93,94,111. My work examining the loading mechanism of the dynein-NudE(L)-Lis1 complex (presented in Chapter
Two), is consistent with previous studies showing that NudE(L) recruits Lis1 to dynein and enhances the AAA+ ring domain-Lis1 interaction that yields the stalled motor 109,110.
142 Unlike the dynein-NudE(L)-Lis1 complex, the mechanism of processivity enhancement observed with the dynein-dynactin motor complex, as compared with dynein alone, remained unclear 12,84. This uncertainty arose from a lack of structural information for the dynein-dynactin supercomplex. Although low- resolution EM structures and biochemical analysis showed the general organization of dynein and dynactin 3,41-46,87,88,112,122,160, the precise location and subunit stoichiometries could not be conclusively determined. Likewise the limited information regarding interactions sites between dynein and dynactin presented little information of how these two large complexes assemble. Our high resolution EM structures of dynein, dynactin, and the dynein-dynactin supercomplex stabilized with the scaffold, BicD2, along with those from the
Carter lab (Discussed in Chapter 3), determined the subunit stoichiometries and organization of these complexes 57,60 and answered the long standing question regarding the structure of the dynein-dynactin supercomplex.
The EM structures of both dynein and dynactin were crucial to solving the first structures of the dynein-dynactin-BicD2 (DDB) complex. Fitting these into the DDB EM density showed that BicD2 facilitates lengthwise interactions between the dynein tail and dynactin’s Arp filament, to provide a binding mode that is distinct from the apparently weak dynein-dynactin interaction mediated by p150Glued and the dynein IC (discussed in Chapter Three). Although this structural work provided many insights into dynein, dynactin and the supercomplex, it also raised questions regarding other coupling scaffolds. An
143
144 Figure 5-1. EM structures of dynein and dynactin. Reconstruction of focused classified negative stained dynein (A) and cartoon (B) showing the subunit organization. Three-dimensional cryo-EM Arp filament structure fitted into the negative stained dynactin envelope (C) and cartoon (B) showing the subdomain and subunit localizations. Cryo-EM structure of the shoulder domain (E) and deconstruction showing the Arp filament proximal (F) and distal (G) three-helix bundles and the β-saddle domain (H). p150Glued (blue), dynamitin (red) and p24
(green) are color coded to indicate which polypeptide is believed to contribute to different domain. The identity of the grey helix remains unclear,
145 important question is whether these interact with dynein and dynactin similarly to
BicD2 to produce similar super-complexes.
Although these structures revealed the precise localization of most dynein and dynactin subunits (Figure 5-1 A-D), the resolution of the dynactin shoulder subdomain was insufficient to identify polypeptide composition, yielding only the general structure (Figure 5-1 C-H) 57,60. My biochemical mapping of protein- protein interactions provides the first model showing how dynamitin, p24 and p150Glued bind to form the interwoven, three-helix bundle shoulder domain structure that is anchored as a unit to the Arp filament (detailed in Chapter Four;
Figure 5-1 E-H). The interactions I defined provide a model for how the p150Glued projecting arm, which binds dynein, may be anchored within the shoulder domain and thus to the Arp filament.
Despite providing a solid understanding of the structure of the dynein and dynactin complexes alone and as a supercomplex, these structural studies fail to answer important questions related to the dynamics of the dynein motor. One is how dynein can switch between the “stalled”, high-force generating dynein-
NudE(L)-Lis1 complex and the processive dynein-dynactin based motor. The fact that the binding sited for NudE(L) and p150Glued on the dynein IC overlap suggests their interactions are mutually exclusive 155, 93,109,110,156. Another area ripe for further investigation is how different “scaffolds” provide additional levels of dynein motor and cargo binding regulation. Furthermore, the mechanisms of scaffold selection, where in the cell, whether on microtubules or in the cytoplasm,
146 and the step-wise assembly process for forming the dynein-dynactin-scaffold supercomplex have yet to be elucidated.
Predicting the structure of the dynein-dynactin-scaffold complexes:
EM structures of the dynein tail-dynactin-BicD2 (TDB) and the microtubule bound dynein-dynactin-BicD2 (DDB) complexes reveal that BicD2 facilitates lengthwise interactions between dynein and dynactin 57,60. BicD2 is just one of a group of putative “scaffolds”, that includes Fip3, Spindly, Hook1 and Hook3. As seen for the DDB complex, dynein-dynactin complexes formed with these scaffolds show enhanced processivity in single molecule motility assays. A common feature is that these proteins contain both an extended coiled-coil and a domain that interacts with vesicular cargos either directly or through additional proteins, such as the Rab family 38,123,184-189. Based upon these features, other proteins that may act as scaffolds are Hook2, RILP, Trak1, Trak2, Golgin160 and
Hap1. Identifying the entire family of coiled-coil “coupling scaffolds” will likely require further screening, potentially including RNAi and/or mutational screens to find candidates that show perturbations to dynein based transport in vivo. These candidates can then be confirmed using established in vitro motility assays.
Defining the family of scaffold proteins is an initial step in understanding the network of dynein motor regulation in vivo, and will be enhanced by elucidating the structures of these complexes. Interaction modeling of the coiled- coil domains of BicD2, Hook1 and Hook3 with dynactin strongly supports that similar interactions stabilize the complex, and they likely generate nearly identical
147 structures (Wenjun Zheng, SUNY Buffalo, unpublished results). Proof of this will require the same EM analysis as was performed for the DDB and TDB structures.
However, these EM studies will only provide a static view of a dynamic process. Additional analysis of how the dynein-dynactin-scaffold (DDS) complex is initiated will determine which subunits are necessary for establishment and maintenance of the complexes. Finally, combinations of in vitro motility assays and in vivo techniques will be needed to elucidate how scaffolds are selected, with wild-type and mutant versions of all components being key to this examination.
Transforming the dynein-Lis1 motor into a motile dynein-dynactin motor complex:
The DDB complex provides the structural archetype for the dynein- dynactin super-complex, but the pathway by which it is assembled has not been defined. Furthermore, the relationship between the “stalled” dynein-NudE(L)-Lis1 complex and the processive dynein-dynactin-scaffold complex is not clear.
Current models favor a view in which the dynein-NudE(L)-Lis1 complex primes the motor for subsequent transformation into the processive motor complex that allows for cargo transport 155 156 96,97,109,110.
Interactions among NudE(L), the dynein IC, and potentially LC8 facilitate
Lis1 binding to one or both of the AAA+ domains, generating a microtubule bound
“stalled” motor 93,94,111,155. Once bound, Lis1 is likely to prevent dynein head
148
149 Figure 5-2. Transformation of the “stalled” dynein-NudE(L)-Lis1 complex into the processive dynein-dynactin motor. (A) The NudE(L) (orange) Lis1 (yellow) bound dynein (blue) complex binds microtubules as a primed motor, whereas dynactin
(green) is targeted to microtubule plus-tips through end-binding proteins. Binding of p150Glued releases the NudE(L)-IC interaction (B) and destabilizes the Lis1-
AAA+ interactions (C) or yields and intermediary complex of dynein, dynactin and
Lis1 (D). This dynein-dynactin interaction is stabilized by interactions with
“coupling scaffolds” (E), which also allows for cargo interactions and processive motility.
150 rotations that allow for the ring stacking observed in the autoinhibited state. This may have the effect of “priming” dynein for motility upon Lis1 release. Consistent with this model, antibody labeling in cultured cells shows dynein and Lis1 localize to the plus-tips of microtubules 152,153, suggesting this may be the site where
“stalled” complexes are formed or accumulate (Figure 5-2 A). This is further supported by data from Aspergillus nidulans, where deletion of endogenous Lis1 drastically reduces the frequency of dynein based vesicle motility events within the hyphal tip, demonstrating that Lis1 is required for initiation of dynein based movements 190. Delivery of dynein and dynactin to the plus-ends of microtubules in A. nidulans required kinesin based transport of vesicles carrying dynein and dynactin towards the microtubule tip.
Transformation of the stalled dynein-Lis1 motor into a processive motor would require release of NudE(L) and Lis1, which likely occurs when p150Glued binds the IC and displaces NudE(L) (Figure 5-2 B). Efficient binding of p150Glued to the IC would require recruitment of dynactin to the plus end of microtubules as well. This may occur via direct microtubule binding of the Cap-Gly and basic domains of p150Glued, but it is more likely to rely on interactions with microtubule end-binding proteins, such as EB1 and Clip170 (Figure 5-2 A) 119,120,191.
Exchange of the NudE(L)-IC interaction for p150Glued binding to the IC may destabilize the Lis1-AAA+ interaction. Alternatively, an intermediary non- processive complex of dynein, dynactin, and Lis1 may be formed (Figure 5-2 C &
D).
151 The final activation step would involve binding of a “coupling scaffold”
(Figure 5-2 E), to lock the motor in a highly processive state. The ring spacing in the DDS complex appears to be optimal for processive motility 96,97, and may differ from the head spacing within the dynein-Lis1 complex. Thus, DDS complex formation might also provide the final push to release Lis1, as the asymmetry imparted on the dynein tail is likely translated into separation of the
AAA+ domains. How scaffolds affect the behavior of the “high-load” generating
Lis1-dynein complex has not yet been evaluated.
The mechanism of scaffold selection is currently a mystery, but may provide insight into the regulation of the dynein motor-cargo interactions in vivo.
The simplest mechanism posits a passive selection where complexes occur by chance when the scaffold is in close proximity to both dynein and dynactin.
Alternatively, the scaffold availability may be regulated either through interactions with cargo or inhibitory interactions with themselves.
Examining the validity of each step of this model will require the combination of in vivo and in vitro assays and necessitate the use of both wild- type and mutant proteins to probe protein-protein interactions and scaffold regulation. Determining how the cargo-carrying DDS motor complex is assembled will be an enormous advancement in understanding regulation of dynein based vesicle transport in vivo.
Concluding remarks
152 It seems as if we have reached a turning point in studying the dynein motor, as many long standing questions are now answered. The field has obtained high-resolution structures of dynein and dynactin, and has shown how these are assembled into a processive motor. Several studies, including my own, have shown how the NudE(L)-Lis1 complex interacts with dynein to generate a
“stalled motor. Now, our focus is turning towards understanding regulatory mechanisms of the dynein motor, which will require integration of structural work, biochemical studies, in vitro motility assays and in vivo imaging techniques to elucidate how the complicated network of regulatory proteins alter dynein functionality.
153
Appendix I
Mycalolide B disrupts the dynactin complex
154 Introduction:
Many sessile organisms produce small molecules that are toxic to other organisms. A general mechanism of toxicity is to alter cytoskeletal dynamics, either by depolymerizing or stabilizing cytoskeletal networks, as seen for
Latrunculins192,193, Cytochalasins194-197, Colchicine198 and taxol199,200. Marine sponges produce several different toxins, known as macrolides, which act to depolymerize actin filaments and sequester G-actin through a common mechanism201-206. These macrolide toxins are composed of a trisoxazole containing macrolide ring and an aliphatic side chain (Figure A1-1 A). Both the ring and side chain can bear a variety of substitutions, with the ring allowing small side group substitutions, such as methyl and hydroxyl groups, and the side chain permitting small or large substitutions.
The actin binding sites of a few macrolide toxins (kabiramide C, jaspisamide A, sphinxolide B, reidispongiolide A and C) have been identified through X-ray crystallography studies201,205. The amino acids the toxin contacts vary slightly, but all bind actin at a site between subdomains 1 and 3 (Figure A1-1
B and C), with the ring likely binding first to the exposed face. Once bound, the aliphatic tail can then intercalate into the cleft between actin monomers and disrupt interactions with subdomain 2 of the adjacent actin protomer, which severs the filament. The macrolide toxin remains bound to the G-actin monomer and prevents its re-incorporation into a filament204. Although the binding sites and mechanisms of action of all macrolide toxins have not been determined, their similar structures suggest they operate via a common mechanism.
155
156
Figure A1-1. Mycalolide B binding site on actin and Arp1. (A) The chemical structure of Mycalolide B 207. (B) Schematic of the actin subdomains, which are conserved in Arp1. The same orientation of this schematic is used for the space filling models in D and E, with the barbed end at the top and pointed end at the bottom. (C) Alignment of bovine actin and Arp1 sequences. Residues responsible for binding the trisoxazole-containing macrolide ring and the side chain are colored yellow and cyan, respectively. Colored bars beneath the alignment correspond with the actin subdomains. (D) Space filling model actin
(PDB-1QZ5) 202, with ATP bound, light green. The residues responsible for binding the ring and the side chain of Mycalolide B are colored yellow and cyan, respectively. (E) Space filling model depicting two Arp1 protomers in a filament
(PDB-5AFT) 57, with ATP bound, light blue. The residues responsible for binding the ring and the side chain of Mycalolide B are colored yellow and cyan, respectively. The amino-terminal 90 amino acids of the dynamitin subunit of dynactin are shown in magenta.
157
Mycalolide B is a partially characterized macrolide toxin, isolated from the
Mycale genus of marine sponges, found in the Bay of Gokasho in Japan. It was first identified as a small molecule inhibitor of actomyosin ATPase activity207.
Initially it was not clear whether the toxin targeted actin or myosin, but it was later found to specifically target and depolymerize F-actin, which led to inhibition of actin-dependent myosin ATPase activity203,204. Because of their ability to induce depolymerization, Mycalolide B and several other macrolide toxins alter the integrity of many cellular structures and impede cell shape changes, adhesion and motility208-212.
Work in neuronal cells, both in culture and in Drosophila larvae, suggests that the inhibitory effects of Mycalolide B are not limited to the actin-based processes. Mycalolide B was found to abolish retrograde transport of dense-core secretory vesicles along microtubules, but did not inhibit anterograde transport213.
This observation is at odds with genetic and antibody inhibition studies, in which altering retrograde motor complexes results in inhibition of microtubule-based transport in both directions 127,214-217. Similar effects were not seen with
Latrunculin A, indicating that the inhibition was unique to Mycalolide B and suggesting that the transport defect was not the result of actin depolymerization and monomer sequestration.
Given that the retrograde transport defect seen with Mycalolide B treatment did not appear to be the result of actin depolymerization, an obvious alternative was that Mycalolide B was acting on the dynactin complex, which contains an actin-like filament comprising eight Arp1 protomers plus a single
158 actin57,60,87,112. Arp1 is >60% identical and >90% similar to actin (Figure A1-1 C)
218, and shares the residues that have been shown to bind other macrolide toxins
(Figure A1-1 D and E) 201,202,205, making it a potential target for Mycalolide B. I explored the possibility that Mycalolide B might disrupt dynactin structure by depolymerizing the Arp1 filament, thus impairing retrograde transport via cytoplasmic dynein.
Results:
Mycalolide B depolymerizes actin and destabilizes dynactin:
I found that Mycalolide B depolymerized actin in a concentration dependent manner, as previously reported 203,204,207. Under the conditions used in this particular experiment, ~50% of the actin could be recovered in the pellet in the absence of Mycalolide B (Figure A1-2 A). Addition of increasing concentrations of Mycalolide B caused depolymerization of the actin filaments, with only a small amount of actin pelleting in the presence of 10 µM and 30 µM
Mycalolide B. Nearly all of the actin appeared to be depolymerized by 60 µM
Mycalolide B. These results differ slightly from the published data, in which the same concentration of actin was completely depolymerized with 3µM Mycalolide
B 203.
After I confirmed Mycalolide B depolymerizes actin, I used sucrose density gradient sedimentation to analyze its effects on F-actin and purified dynactin. As seen in the previous assay, DMSO did not alter the polymerization state of actin as pelletable F-actin could be detected in the presence of DMSO (Figure A1-2 B,
159
160 Figure A1-2. Mycalolide B depolymerizes actin and disrupts the dynactin complex.(A) Actin pelleting assays show that the presence of DMSO does not alter the polymerization of actin as compared with general actin buffer (GAB), while the addition of Mycalolide B causes depolymerization of actin as determined by the amount of actin in the supernatant (S) vs. the pellet (P). (B)
Sedimentation of actin through a 5-20% sucrose (w/v) gradient in general actin buffer. In the presence of DMSO a fraction of this actin sample pelleted completely through the gradient, while Mycalolide B eliminates this pellets. (C)
Sedimentation of bovine dynactin through 5-20% sucrose (w/v) gradients. The
DMSO control gradient shows that the dynactin subunits are present in the pellet, owing to the very low ionic strength of the buffer. The addition of varying concentrations of Mycalolide B alters the sedimentation behavior of the dynactin subunits such that they no longer pellet, indicating dynactin has been disrupted into subcomplexes and monomeric proteins.
161 top). Addition of 30 µM Mycalolide B caused this pool to depolymerize completely (Figure A1-2 B, bottom).
The same sedimentation assay was used to determine the effect of
Mycalolide B on dynactin. Under control (i.e. DMSO) conditions, purified dynactin pelleted through the sucrose gradient, due to the very low ionic strength of the buffer (Figure A1-2 C, top left). Mycalolide B caused disruption of the dynactin complex in a concentration-dependent manner (Figure A1-2 C, top right and bottom). Dynactin subunits were present in the fractions across the gradients. Some appear to cosediment, suggesting the presence of subcomplexes as is seen when the Arp1 filament is depolymerized using the chaotropic salt, potassium iodide (KI) 87.
I pursued this possibility using sucrose gradient sedimentation under conditions of higher ionic strength where dynactin would not pellet (DMSO control; Figure A1-3 A top). In the presence of Mycalolide B, instead of cosedimenting as a large particle, dynactin subunits showed distinct sedimentation behavior. The Arps sedimented as a tight peak in the light sucrose fractions, consistent with complete disassembly of the filament (Figure
A1-3 A, bottom). p150/p135, dynamitin and p24 appeared to cosediment (Figure
A1-3 A, bottom), reminiscent of what is seen following KI disruption, suggesting that the shoulder/arm complex is released as a distinct entity following filament depolymerization. The pointed end complex subunits (Arp11, p62, p27 and p25) could not be detected via Coomassie blue staining.
162 Mycalolide B disrupts the dynactin complex in cytosol:
I next examined whether Mycalolide B had the same effect on dynactin in a detergent lysate prepared from cells as it does on purified dynactin.
Sedimentation of a detergent lysate of HEK293T cells showed that dynactin subunits sediment at 20 S along with dynein, as indicated by the presence of dynein intermediate chain (Figure A1-3 B, top). Treatment of the HEK293T lysate with Mycalolide B once again disrupted the dynactin complex, causing the individual subunits to sediment at distinct places in the sucrose gradient. Dynein
(intermediate chain) still sedimented at 20S, indicating the disruptive effect of
Mycalolide B is selective (Figure A1-3 B, bottom). Densitometric analysis revealed that 60.9% of the Arp1 was present in light sucrose fractions in the
Mycalolide B treated sample versus 4.4% of in the control, indicating significant disassembly (Figure A1-3 C). The incomplete Arp filament disassembly seen in the cytosol sample is most likely due to the presence of actin. The large pool of actin presumably acts to sequester the Mycalolide B, reducing its effective concentration and resulting in incomplete depolymerization of dynactin’s Arp1 filament.
Discussion:
Mycalolide B depolymerizes actin filaments, and I showed it is also able to depolymerize Arp filaments. Given that actin and Arp1 have highly similar structures, the mechanism of severing the Arp1 filament is likely the same as for actin filament severing. The trisoxazole ring likely binds to the exposed face of
163
164 Figure A1-3. Mycalolide B disruption of dynactin in vitro and in vivo.
(A) Analysis of Mycalolide B on dynactin integrity in vitro. Purified bovine dynactin treated with DMSO or 2 µM Mycalolide B was subjected to sedimentation into a
5-20% sucrose gradient. Sedimentation indicates that the Arp minifilament is fully depolymerized into a monomer/dimer pool, while the shoulder/sidearm subunits cosediment similar to potassium iodide disruption. (B) Analysis of MB on dynactin integrity in vivo. HEK293T cell lysate treated with DMSO or 2 µM
Mycalolide B was subjected to sedimentation into 5-20% sucrose gradients.
Immunoblots indicate that Mycalolide B does not cause a full disruption of the
Arp minifilament. (C) Densitometry indicates about half of the Arp1 in a monomer/dimer pool in the presence of Mycalolide B vs. <5% in the vehicle control. The different extent of Arp1 disassembly seen with purified dynactin vs. dynactin in cytosol is due to the presence of actin and other cytosolic components.
165 subdomains 1 and 3 and the aliphatic tail inserts into the cleft formed between subdomain 1 and 3 and subdomain 2 of the adjacent Arp1 monomer.
Interestingly, the site the trisoxazole ring binds on Arp1 is partially blocked by a
α-helix belonging to the amino-terminus of dynamitin (Figure A1-1 E). This suggests a couple of mechanisms by which Mycalolide B may disrupt dynactin.
One possibility is that Mycalolide B does not bind Arp1, but instead binds the conventional actin monomer found at the pointed end of the Arp filament. By binding conventional actin, Mycalolide B would sever the pointed end complex from the Arp filament. Experiments in cultured cells using siRNAs targeted to the pointed end complex subunits p62 and Arp11 indicate that loss of these components destabilize the Arp filament and lead to release of shoulder/arm components.
Alternatively, Mycalolide B may bind at multiple points on the Arp filament to sever it. In addition to this severing, the Mycalolide B binding site on Arp1 overlaps with that of dynamitin, so binding of Mycalolide B might displace the dynamitin N-terminus from the Arp1 filament, resulting in a release of the shoulder/arm as an intact subcomplex, similar to what is observed with dynamitin overexpression.
I propose that Mycalolide B acts as a combination of these mechanisms.
Given that we know that the pointed end complex is needed for the stability of the
Arp filament, once it has been removed, the filament would be destabilized.
Additionally, displacement of dynamitin and the associated shoulder/arm components (p150Glued and p24) would inhibit the dynein-dynactin-cargo
166 interaction, effectively inhibiting retrograde transport, even without full depolymerization of the Arp filament. This may explain why retrograde transport can be fully abolished with 2 µM Mycalolide B, which does not completely depolymerize of the Arp filament in HEK293T lysates.
It is possible that the free Arp1 produced by Mycalolide B treatment can reassemble into filaments. Without dynamitin to regulate the length of the Arp1 filament, any Arp1 not bound to Mycalolide B would assemble into untemplated filaments, explaining the broad distribution of Arp1 across the sucrose gradients
(Figure A1-3 B, bottom).
Although Mycalolide B disrupts the dynactin complex and effectively abolishes retrograde transport when applied to D. melanogaster nerve terminals and cultured hippocampal neurons, it seems unlikely that it will be widely useful for in vivo studies due to the side effects of actin depolymerization. Despite this,
Mycalolide B and likely other macrolide toxins will have utility in in vitro motility studies as a fast acting inhibitor of the dynein-dynactin motor.
Materials and Methods:
Purification of dynein and dynactin:
Dynein and dynactin were purified from bovine brain as described previously (Jim Bingham) with the following changes. Dynein and dynactin were eluted from the MonoQ 10/100 column using a linear salt gradient. Dynein and dynactin containing fractions were identified by Coomassie blue stained SDS polyacrylamide gels, then rechromatographed separately on a MonoQ 5/50
167 column. Elution from the MonoQ 5/50 column was performed with a salt bump, the concentration of salt was determined from the conductivity measurements of the first and last fractions pooled (described in Chapter 3).
HEK293T cell culture:
HEK293T cells were grown in high glucose DMEM (Gibco), with 10% FBS and 1% L-Glutamine at 37°C with 5% CO2. Detergent lysates were prepared by washing the cells with PBS, then incubating at room temperature in 2 ml of
0.25% Trypsin-EDTA (Gibco) for 2 minutes. After incubation, 8 ml of DMEM, with 10% FBS and 1% L-Glutamine was added and pipetted to resuspend the cells. This cell suspension was pelleted at 50 in a dynac centrifuge. The pellet was resuspended in 1 ml of cold PBS and pellet at 2,100 RFC at 4°C for 10 minutes. The pellet was then resuspended in lysis buffer, 20 mM Tris pH 7.4,
150 mM NaCl, 0.1% Triton X-100, 1 mM EGTA, 2 mM β-ME, with protease inhibitors, and incubated with gentle agitation for 1 hour. The lysed sample was then pelleted at 20,800 RCF for 10 minutes at 4°C. The supernatant was used within 1 hour for sedimentation assays.
Actin polymerization:
Lyophilized human platelet actin (Cytoskeleton) was resuspended as a
10X solution at a concentration of 10 mg/ml in MilliQ water, as per instructions.
The actin stock (25 µl) was diluted with general actin buffer (225 µl), 5 mM Tris,
0.2 mM CaCl2, 0.2 mM ATP pH 8.0, and incubated on ice for 30 minutes. A volume of actin polymerization buffer, 500 mM KCl, 20 mM MgCl2, 10 mM ATP
168 pH 7.5, equal to one tenth of the actin solution was added (25 µl), the solution was incubated at room temperature for 2.5 hour to allow for polymerization.
After polymerization 15 µl of general actin buffer and 10 µl DMSO or Mycalolide
B (Wako) stocks were added to the actin solution to yield final concentrations of
60 µM, 30 µM, 10 µM and 0 µM Mycalolide B then incubated at room temperature for 30 minutes. Additionally a sample with 25 µl of general actin buffer was assayed to confirm that DMSO did not effect actin polymerization.
Actin pelleting was performed using an SW55 rotor, at 40,400 RPM for 1.5 hours, 24° C. After pelleting, the supernatant was removed and the pellet was resuspended in 400 ul of SDS gel loading buffer, 400mM Tris pH 8.8, 0.25 mM β-
ME, 10% glycerol, 1% SDS, 0.01% BPB, 0.005% Pyronin Y, and boiled immediately. 12 µl of the supernatant and 16 µl of the pellet were run on an 8%
SDS polyacrylamide gel and Coomassie stained to analyze the polymerization of the actin.
Sucrose density gradient analysis:
50 µg of pre-polymerized actin or purified bovine dynactin diluted in 285 µl of general actin buffer were incubated with DMSO or Mycalolide B, 5% of final volume, for 30 minutes. The samples were layered onto 4.7 ml 5-20% sucrose
(w/v) gradients in general actin buffer and centrifuged 29.9 K RPM for 16 hours at 24° C. Ten 0.5 ml fractions were collected from each gradient, proteins that sedimented through the gradient were resuspended in 50 µl of SDS gel loading buffer. 100 µl of each fraction were subjected to TCA precipitation then run,
169 along with 10 µl of the pellet, on 12.5% SDS polyacrylamide gels and Coomassie blue stained to determine the sedimentation behavior.
Alternatively, 50 µg of purified bovine dynactin or 250-500 µg of a
HEK293T cell detergent lysate was incubated for 30 minutes at room temperature with either DMSO (2% of final volume) or 2 µM MB (in DMSO) in a final volume of 300-500 µl in 35 mM Tris, 5 mM MgSO4, pH 7.2 (TM buffer). The samples were then layered onto 11.8 ml 5-20% sucrose (w/v) gradients in TM buffer and centrifuged for 17.5 hours at 34K RPM in an SW41 rotor. Twelve 1 ml fractions were collected from each gradient. For the purified dynactin samples,
100 µl of each fraction was subjected to TCA precipitation and run on a 12.5%
SDS polyacrylamide gel. For the HEK293T lysate samples, the entirety of each fraction was subjected to TCA precipitation, then run on a 12.5% SDS polyacrylamide gel and transferred to PVDF membrane for immunoblotting. The following antibodies were used to detect dynactin subunits: mouse monoclonal antibodies 150B and 45A were produced and purified as previously described, mouse anti-dynamitin (BD Biosciences Pharmingen), and rabbit anti-DCTN6
(Proteintech Group). The mouse monoclonal antibody 74.1 (Millipore) was used to detect dynein intermediate chain as a sedimentation standard and internal control.
170
Appendix II
Analysis of recombinant p150Glued fragments
171
Introduction:
My initial motivation for reconstituting the dynactin shoulder complex from bacterially expressed proteins was to generate large amounts of a dynactin subcomplex that could interact with a complementary complex from dynein
113,114,117,155. I hoped that this recombinant subcomplex could be used in structural analysis to generate a dynein-shoulder super-complex, allowing us to get a first sense of how the dynein-dynactin super-complex forms. At the time I initiated my work, only the interaction between p150Glued AA 415-530 and the N- terminus of the dynein intermediate chain (IC, AA 1-32), had been identified
113,114,117. Evidence that this interaction is relevant to dynein based transport is abundant: overexpression of p150Glued AA 217-548 or injection of antibodies that bind the IC N-terminus inhibit dynein-based motility in cultured cells 113,165 and an antibody that binds p150Glued AA 217-548 prevents dynactin from stimulating dynein-driven bead movements in vitro 167. These data suggest that the direct interaction between p150Glued and DIC is required for dynein-dynactin based motility in vivo and in vitro. We now know that formation of a highly processive dynein-dynactin complex involves extensive lengthwise interactions between dynein, a “coupling scaffold”, such as BicD2 (Discussed in Chapter 3), and the
Arp filament domain of dynactin 57,60.
Single particle EM on platinum coated or negatively stained samples of dynactin reveal a 24 nm long projecting arm terminating in a globular mass
(Figure A2-1 A)112,160. Decorating with the monoclonal antibody 150.1, which
172 recognizes p150Glued AA 217-548, showed the globular domain to be labeled 112, indicating that the projecting arm includes p150Glued AA 1-548. Furthermore, subtraction of the mass of the Arp filament from a low-resolution three- dimensional EM structure (Figure A2-1 B), suggests that the shoulder domain contributes 357-418 kDa in mass (Figure 2-1 C). Assuming the shoulder incorporates the entirety of dynamitin and p24, this mass requires an additional
≈129-190 kDa to be accounted for. This remaining mass can only be attributed to p150Glued, which I calculated to correspond with a dimer of ≈590-860 amino acids, which is consistent with the projecting arm comprising the N-terminal ≈550 amino acids.
This structural information, combined with our knowledge that we could generate recombinant complexes of dynamitin and p24, with 2:1 stoichiometry that matches native dynactin 87,178, provided the basis for my work. I took advantage of the established protocol where insoluble bacterial expressed p24 is combined with soluble dynamitin under denaturing conditions and dialyzed out of the denaturant to form a soluble complex of dynamitin and p24, thus “rescuing” p24 solubility with a native binding partner 178. My goal was to identify a p150Glued fragment capable of binding the dynein IC that could be incorporated with dynamitin and p24 into a recombinant shoulder/arm complex. Although ultimately I was not able to reconstitute a p150Glued/dynamitin/p24 complex, my efforts yielded important new insights on subunit organization in the shoulder domain and the mechanism of p150Glued anchoring (Chapter Four). In this
173
174 Figure A2-1. Dynactin structure. (A) Platinum coated, freeze etched dynactin particle with the Arp filament and projecting arm labeled. (B) 34Å resolution three-dimensional reconstruction of dynactin with fitted Arp filament, modeled with six Arp1 protomers. (D) The resulting shoulder domain mass after subtraction of the Arp filament from the three-dimensional EM density.
175 Appendix, I will describe my efforts to isolate soluble p150Glued fragments that we now know comprise the projecting arm, rather than the shoulder.
Results: p150Glued fragment design:
The first p150Glued fragment I designed contained AA 217-1048. My hope was that inclusion of the two readily soluble coiled-coil domains, AA 217-548 and
AA 926-1048 165 113, would enhance solubility of the intractable central region (AA
549-925). This turned out not to be the case, as the fragment was insoluble when expressed in bacteria. However, the AA 217-1048 fragment could be refolded by dialysis out of the denaturant Guanidine HCl, as performed for dynamitin and p24 178. The resulting soluble protein appeared to be a large oligomeric complex when analyzed using Size Exclusion Chromatography (SEC)
(Table A2-1). Given this oligomerization, I attempted to renature this p150Glued fragment in the presence of both dynamitin and p24, hoping that either, or both, would bind and produce a complex resembling the native shoulder/arm domain.
This was based in the observation that recombinant dynamitin forms oligomeric complexes, but only forms trimers in the presence of p24 178, thus, I reasoned that this might also occur with p150Glued. Although p24 solubility was rescued with p150Glued AA 217-1048, the resulting complex eluted in the column void when analyzed with SEC, suggesting that a large oligomeric complex had still formed (Table A2-1). I was unable to detect any interactions between p150Glued
AA 217-1048 and dynamitin.
176 Given this, I set out to design additional p150Glued fragments that might be better behaved. To do so I used prediction programs to identify boundaries between secondary or tertiary structures (Coils, DisEMBL, MARCOIL, Paircoil2 and PSIPRED) as well as sequence conservation analysis to reveal regions that might play key roles in protein-protein interactions. Secondary structure predictions indicated that p150Glued is largely α-helical, with AA 217-548, AA 926-
1048 and AA 1095-1103 showing coiled-coil propensity. The first two coiled-coil domains are well conserved (67.2% and 66.9% identical, respectively), as is the intervening region, which I broke into three regions, AA 537-629, AA 630-735 and AA 736-931 (65.8%, 61.5% and 57.2% identical, respectively) (Figure A2-2
A). I subcloned these into bacterial expression vectors. Some were appended to the entirety or portions of adjacent coiled-coil domains in the hope that this would favor solubility (Figure A2-2 B).
Solubility analysis of p150Glued fragments:
As expected, the coiled-coil domains, AA 217-554 and AA 932-1048, were soluble, as were smaller fragments of the first coiled-coil domain, AA 217-420,
AA 421-536, AA 421-554 and AA 217-536 (Table A2-1). However, inclusion of any portion of AA 555-931 interfered with solubility.
I then used the renaturation method for generating recombinant dynamitin- p24 trimers to try to produce soluble p150Glued fragments 178. Briefly, the insoluble protein is purified under denaturing conditions, then renatured using step-wise dialysis out of denaturant. Attempts to resolubilize the p150Glued
177
178 Figure A2-2. Structural prediction of p150Glued domains used to design recombinant fragments. (A) ClustalOmega alignment of p150Glued sequences, color coded for percent identity. Predicted secondary structures (top) and coiled- coil/multi-coil propensity (bottom) for rat p150Glued, breaks in these correspond with sequence insertions in the alignment. (B) Schematic of the p150Glued secondary structure with the coiled-coil and intercoil-domain labeled and the recombinant expression constructs generated. The fragments in grey were not analyzed.
179
180 Table A2-1. Analysis of p150Glued fragments indicating fragment solubility and properties after renaturing alone or with p24. The behavior of full-length p24 and p24 AA 1-67 were identical in this analysis.
181 fragments by themselves yielded soluble protein, but SEC analysis indicated that almost all of the p150Glued fragments were large, oligomeric complexes that elute in the column void (Table A2-1). The exceptions to this were the previously characterized coiled-coil fragments, AA 217-554 and AA 932-1048, which eluted with Stokes’ radii of 62.8 ± 0.2 Å and 42.0 ± 2.0 Å, respectively. An additional fragment, AA 736-1048, showed some promise, as a small portion of this fragment eluted with a Stokes’ radius of 54.9 ± 0.1 Å, with the majority eluting in the column void.
Since all of the new fragments were soluble as oligomeric complexes on their own, I tried renaturing the p150Glued fragments in the presence of p24 in hopes that p24 may reduce the propensity of the p150Glued fragments to oligomerize. To perform this analysis I used either full-length p24 or p24 AA 1-67, which contains the first α-helical domain, AA 1-32, that I previously showed did not interact with dynamitin (discussed in Chapter Four). The initial work with the
AA 217-1048 fragment showed that it yielded soluble complexes with full-length p24 and AA 1-67 (Table A2-1). As p24 could not be renatured with either of the coiled-coil domains, AA 217-554 or AA 932-1048 (Table A2-1), this led me to conclude that the p150Glued-p24 interaction might involve to AA 555-931 (Table
A2-1). However, the observation that the soluble p24-p150Glued complexes eluted as large oligomers in the column void, suggested that the rescue of p24 solubility might be due to non-specific protein-protein interactions, with the p150Glued oligomers trapping the p24 polypeptide, causing them to appear soluble.
182 I pursued this hypothesis by trying to identify smaller fragments of p150Glued that were sufficient to form a soluble complex with p24 that did not elute in the column void. Although all of the fragments tested (AA217-629, AA 217-
735, AA 421-735, AA 537-1048, AA 555-932, AA 630-1048, and AA 736-1048) formed soluble complexes with full-length p24 and p24 AA 1-67, all eluted in the column void when analyzed with SEC (Table A2-1). These results support the conclusion that the apparent p24 solubility is the result of non-specific protein- protein interactions, likely protein trapping by the oligomers. Had p24 only interacted with a subset of overlapping fragments, it would have suggested a region of p150Glued responsible for binding p24 within the dynactin shoulder/arm.
Discussion:
Analysis of solubility of p150Glued fragments expressed in bacteria indicates that AA 555-932 interferes with solubility. This is most likely because this region is unable to properly fold into its native structure without eukaryotic chaperones.
Prediction programs indicate that this region is largely α-helical, with some portions being hydrophobic, consistent with a globular protein (Figure A2-3).
Initially I believed that this region of p150Glued was buried within the shoulder domain and required additional protein-protein interactions with dynamitin and p24 to fold properly, thus explaining why it was insoluble when expressed in bacteria by itself. However, recent EM work shows that this domain is globular and solvent exposed 57, suggesting that insolubility of the recombinant p150Glued fragments arises from misfolding rather than the absence of native binding
183 partners. Furthermore, proteomics analysis indicates that multiple residues between AA 555-932 are phosphorylated, ubiquitinated, and acetylated in vivo
(PhosphoSitePlus, MaxQuant Database and SwissPalm), supporting the idea that post-translational modifications may contribute to proper folding and solubility.
Furthermore, recent EM work indicates that p150Glued AA 1048-1281 is the portion that incorporates into the shoulder 57. Although we assumed this part of p150Glued contributed to the shoulder domain, it was not included in the fragments generated as previous work in the lab showed that it was highly sensitive to proteolysis (Frances Cheong, Ph.D. Thesis). Instead, the fragment I used, AA
217-1048, is now known to be the projecting arm 57 which does not interact with either dynamitin or p24. This is consistent with my findings that p150Glued AA
217-1048 did not interact with any dynamitin fragments and that the copurification of p150Glued with p24 after renaturation was the result of non- specific protein-protein interactions.
Although my biochemical work with p150Glued failed to provide insight into the interactions among p150Glued, dynamitin and p24, structural predictions combined with the recent EM work allows for a model of how p150Glued incorporates into the shoulder (discussed in more detail in Chapter 4). The C- terminus of p150Glued contains three predicted β-strands, AA 1172-1178, AA
1250-1257 and AA 1265-1270, which likely account for the six β-strands in the dimerization domain, as two copies of p150Glued are present in dynactin. The four
α-helical domains that underlie these β-strands are approximately 25 amino
184 acids in length, and likely correspond with the first α-helical domain of p24 AA 5-
22 and p150Glued AA 1190-1217. Additionally, as the p150Glued EM density can be traced within the shoulder, nearly connecting with an α-helix in the hook domain, it is likely that it also interacts with dynamitin. This three-helix bundle is believed to correspond with AA 145-195 of dynamitin and p150Glued AA 1095-
1130. These interactions weave the p150Glued C-terminus into the shoulder, anchoring the projecting arm to the remainder of dynactin.
Although generating a recombinant shoulder domain that might be useful for high-resolution structural studies will not require AA 555-932, producing a recombinant dynactin holo-complex or shoulder/arm complex that can bind the dynein IC will require a system where this region can be expressed in soluble form. The best candidate for expression is the Sf9 system which has been used to successfully express recombinant dynein complexes. However, the combination of post-translational modifications and the complicated stoichiometry of dynactin subunits may hinder this. Regardless, recombinant dynactin would be an invaluable tool as it would provide a way to prepare disease-associated dynactin mutants that could be used for functional studies in vitro.
Materials and Methods
Cloning
Fragments of p24 were cloned using the full-length pRSET-p24 vector as a template (described in Chapter 4) 178. p150Glued fragments were generated using full-length rat p150Glued (p60 DNA, gift from Erika Holzbaur) using PCR
185 amplification with the appropriate primer combination (Table 1). The amplicon was ligated into pRSET-A using EcoRI and XhoI restriction enzymes.
Protein purification:
Fragments of p150Glued were expressed and purified using a modified protocol for full length p24 178. Bacterial cultures for the fragments were grown at
37°C to an OD600 of 3-4, induced with 10 mM IPTG then grown at 37°C overnight.
After centrifugation, the pellet was resuspended in 100 ml resuspension buffer,
100 mM Tris, 5 mM EDTA, 5 mM DTT, pH 7.0, per liter of bacterial culture, then lysed using an Avestin EmulsiFlex C3. After lysis, the insoluble material was separated from the supernatant with centrifugation. The insoluble pellet was washed with 20 ml of wash buffer, 100 mM Tris, 2 M Urea, 5 mM EDTA, 5 mM
DTT, 2% Triton X-100, per liter of bacterial culture, then centrifuged to separate the insoluble and soluble material. This wash step was repeated an additional four times, the final pellet was rinsed with the resuspension buffer to remove detergent, then resuspended in 20 ml of 6 M Guanidine-HCl, 0.5 M NaCl, 20 mM
Sodium Phosphate at pH 7.4. This was loaded onto a 5 ml His-Trap (GE Life
Sciences), washed until the baseline returned to zero, then eluted with a pH gradient from pH 7.4 to pH 2.1. Protein purity was assessed with Coomassie
Blue stained SDS-PAGE.
186 Refolding complexes:
Fragments of p150Glued or p150Glued-p24 complexes were renatured as previously described 178. Briefly, p150Glued and p24 fragments were dialyzed against 3 M Guanidine-HCl, 0.5 M Arginine, 20 mM Tris, pH 7.4 at 4°C, then into
2 M Guanidine-HCl, 0.5 M Arginine, 20 mM Tris, pH 7.4 at 4°C individually. After dialysis, the p24 and p150Glued fragment were combined with 1:1 stoichiometry, dialyzed further into 1 M Guanidine-HCl, 0.5 M Arginine, 20 mM Tris, pH 7.4 at
4°C, then 0.5 M Arginine, 20 mM Tris, with 1 mM DTT, pH 7.4 at 4°C. The final dialysis buffer contained 20 mM Tris, 100 mM NaCl, 1 mM DTT, pH 7.4 at 4°C.
After the final dialysis step, the p150Glued or p150Glued-p24 sample was centrifuged at 14 K RPM in an Eppendorf Centrifuge 5417C for 20 min at 4°C to remove insoluble protein. The resulting supernatant and pellet were analyzed with Coomassie Blue stained SDS-polyacrylamide gels to determine whether the fragment or complex was soluble or insoluble after renaturation.
Size exclusion chromatography analysis
Purified p150Glued -p24 complexes were loaded onto a pre-equilibrated
Superose 12 HR 10/20 (GE Life Sciences) with 20 mM Tris, 150 mM NaCl, 1 mM
β-Mercaptoethanol, pH 7.4. A 0.5 ml protein sample was injected into the column at 0.2 ml/min, the column continued to run at 0.2 ml/min with 0.5 ml fractions collected throughout the entirety of the column run. Stokes’ radii were calculated using a standard curve generated with standards from both the Gel
Filtration HMW and LMW Calibration kits (GE Life Sciences) according to the
187 protocol for the Superose 12 HR 10/30 column. Fractions from the included volume of the Superose 12 HR 10/30 column were analyzed with Coomassie
Blue stained SDS-PAGE.
188
GGAG GAAC TGTGG AGATGC GAGGGAC GCTCAAG GGATGCT CCTGGCT CCTGGGC TCAGCTCC AGGTCTCTGG TGCTGAGACC GTGCTGCAGG CCTCAATAGTGCG
Sequence CCGCTCGAGGACCTGAGTGA CCGCTCGAGGAGCAGCCCGA CCGCTCGAGCAGGAAGCGTC CCGCTCGAGGCAGAGATCAC CCGGAATTCTCAAAACTTCT CCGGAATTCTCAGCTCTCTG CCGGAATTCTCAAGCGAGGT CCGCTCGAGACCATCGACGA CCGGAATTCTCACTGGTTGG CCGCTCGAGGAGGAAGGGCT CCGCTCGAGAAAATCAAGTT CCGGAATTCTCAGAAGTCGA CCGGAATTCTCATGCACGCA CCGGAATTCTCACCGGAGTC
AA 217AA forward 421AA forward 537AA forward 555AA forward 630AA forward 736AA forward 932AA forward AA 420AA reverse 536AA reverse 554AA reverse 629AA reverse 735AA reverse 932AA reverse AA 1048AA reverse PrimerName Glued Glued Glued Glued Glued Glued Glued Glued Glued Glued Glued Glued Glued Glued p150 p150 p150 p150 p150 p150 p150 p150 p150 p150 p150 p150 p150 p150
1 2 3 4 5 6 7 8 9 10 11 12 13 14 No.
189 Table A2-2. PCR primers used to generate p150Glued fragments. Combinations of these (forward and reverse) were used to generate the various p150Glued fragments that were ligated into pRSET-A after digestion with EcoRI and XhoI.
190
Appendix III
The p27 free pool is Rab11a sensitive
191 Introduction:
Early investigation of the structure and functions of the dynactin complex utilized monoclonal antibodies generated against individual subunits. Some were used for immunogold labeling of cells and single particle EM analysis, whereas others were used to determine dynactin localization via light microscopy. One antibody used for immunofluorescence was mAb 27A, a monoclonal antibody obtained in a screen of hybridomas obtained from mice immunized with purified bovine pointed end complex 177. Sucrose gradient sedimentation of detergent cell lysates showed that most dynactin subunits are present only as part of a ≈20
S complex, whereas the p27/p25 heterodimer is in both the lighter sucrose fractions and the ≈20 S fractions, indicating a subset of molecules that are not bound to dynactin (Nicholas Quintyne, Ph.D. Thesis; Brett Scipioni, Ph.D. Thesis).
Because mAb 27A was able to detect the free p27 pool, we reasoned it would be a useful probe to investigate the behavior of this pool in whole cells.
Immunofluorescence with mAb 27A revealed two conspicuous patterns: dispersed puncta and a perinuclear cluster of puncta, in addition to the centrosomes staining expected for intact dynactin 32,64,165,174,219,220 (Figure A3-1 A
& B). Quantification indicated that ≈55% of cells had a cluster and the remaining
≈45% of cells had either dispersed puncta or diffuse hazy staining (Nicholas
Quintyne, Ph.D. Thesis). These distinct localizations suggested to us that p27/p25 might show cell cycle dependent behavior. This was confirmed using synchronized cells. Cells in G1 had a diffuse or hazy staining pattern. As the
192
193 Figure A3-1. Monoclonal antibody 27A staining is cell cycle dependent.
Representative images of Cos7 cells stained with the mAb 27A, showing a perinuclear cluster (A) and dispersed puncta (B). (C) Cartoon depiction of the cell cycle dependent localization of p27 (Nicholas Quintyne, Ph.D. Thesis). In G1 mAb 27A staining shows a hazy background throughout the cytoplasm. As cells enter S-phase the staining becomes more punctate and clusters near the nucleus during G2.
194 cells entered S-phase dispersed puncta became more predominant. These accumulated in the perinuclear region as the cells entered G2 (Figure A3-1 C)
(Nicholas Quintyne, Ph.D. Thesis).
The cell-cycle dependent localization seen with mAb 27A was not observed with antibodies against any other dynactin subunits. Therefore, we set out to determine the identity of the labeled structures and examine potential regulation. Phosphorylation was an obvious candidate, as two-dimensional gel electrophoresis showed that p27 is phosphorylated during M-phase, at a site later determined to be threonine 186 172. However, labeling cells overexpressing phospho-mimic or null mutants (T186D and T186A, respectively) with mAb 27A yielded identical centrosomal staining patterns, suggesting that phosphorylation was not the basis for the cell cycle dependent change in localization and further indicating that endogenous p27 behaves differently from exogenously expressed forms (Brett Scipioni, Ph.D. Thesis).
Operating under the assumption that the labeled structures were endomembranes, I tried to gain insight into the identity of the compartment using
Rab GTPases, which are known to associate with different membranous organelles and regulate vesicular trafficking (reviewed in 221). I used overexpression of different Rab GTPases to alter intracellular compartment dynamics, hoping also to alter the localization of the immunostained compartment. In a limited screen using overexpression of wild type Rab
GTPases (Rab 1a, 5, 6a, 7, 9 and 11a) none were found o affect the structures labeled by the mAb 27A (data not shown).
195 Another Rab, Rab11a, associates primarily with the recycling endosome and regulates trafficking of molecules within the slow recycling pathway for which the transferrin molecule is a canonical marker 222-224. Overexpression of wild- type or constitutively active (Q70L or S20V) Rab11a does not alter the delivery of transferrin to the perinuclear recycling endosome, whereas overexpression of the dominant negative mutant (S25N) shows a reduction of transferrin in the perinuclear region 224-227. Although trafficking of other cargos can be altered by
Rab11a overexpression, I chose to use transferrin as a probe to confirm the behavior of the Rab11a mutants before testing their effect on mAb27A localization.
Results:
Transferrin receptor trafficking in cells overexpressing Rab11a mutants:
Untransfected control cells or those overexpressing the Rab11a constructs were continuously labeled with Alexa-fluor 488 or 546 labeled transferrin, then chased with media lacking labeled transferrin. Cells were fixed at 20 min and 60 min during the continuous labeling and also after a 60 min chase period. Analysis shows that the untransfected control cells have a visible perinuclear cluster of transferrin after 20 min, which becomes more prominent at the 60 min time point. This fluorescence intensity decreases dramatically and the staining pattern becomes highly dispersed after the 60 min chase, indicating that transferrin has been recycled to the plasma membrane where it is released into the media (Figure A3-2 B). Cells overexpressing Rab11a wild-type or the
196
197 Figure A3-2. A dominant negative Rab11a mutant alters transferrin localization and trafficking. (A) Time course of the Alexa-488 or 546 labeled transferrin uptake assay used to monitor transferrin behavior. Cos7 cells were transfected with GFP (not shown) or mCherry (A) controls or Rab11a wild-type (B & C), constitutively active (CA; D & E) or dominant negative (DN; F & G), fixed at the indicated time points and imaged to determine how transferrin labeling was affected by the different experimental treatments.
198 constitutively active mutants (Q70L and S20V) show the same behavior as controls (Figure A3-2 C-F). By contrast, cells expressing the dominant negative
Rab11a mutant (S25N) show normal transferrin uptake, but the probe fails to localize to the perinuclear cluster, exhibiting delayed accumulation and washout
(Figure A3-2 G and H).
Rab11a overexpression alters the localization of the p27 cluster:
Once I had confirmed that overexpression of the Rab11a constructs yielded the expected phenotypes, I examined how they altered the localization of the mAb 27A stained structures. Cells overexpressing the parent plasmid (GFP or mCherry vector) or wild-type Rab11a showed normal mAb 27A labeling in asynchronous cells, of which ≈55% show with a clustered pattern (Figure A3-3).
Overexpression of either the constitutively active or dominant negative Rab11a mutants altered localization, having opposite effects on the mAb 27A labeling pattern. The constitutively active Rab11a mutants (Q70L and S20V) result in a more dispersed staining pattern (≈77% of cells), whereas the dominant negative
Rab11a mutants (S25N) caused the stained structures to remain clustered
(≈76% of cells) (Figure A3-3). This suggested that the mAb 27A labeled compartment associates with a dynamic membrane compartment regulated by
Rab11a.
199
200 Figure A3-3. p27 localization is altered in Cos7 cells overexpressing Rab11a mutants. Asynchronous Cos7 cells overexpressing mCherry or GFP control plasmids or Rab11a wild-type (WT) show normal p27 distribution patterns, with
≈55% showing a p27 cluster. Overexpression of constitutively active Rab11a
(S20V or Q70L) causes a decrease in the number of cells with a p27 cluster
(≈23%), whereas overexpression of the dominant negative Rab11a (S25N) increases the number of cells with a p27 cluster (≈76%).
201 Discussion:
Although I was unable to determine the identity of the compartment labeled with mAb 27A, my data indicate that it is Rab11a sensitive, and most likely a recycling endosome. The cell cycle dependent localization of this compartment suggests it may play an important role during different stages of the cell cycle, when specific membrane compartments are targeted for dissociation and transport. One possibility is that it helps to prepare the cell for mitosis and cytokinesis, when recycling endosomes are trafficked to the cleavage furrow to provide additional membrane allowing for the final steps of cytokinesis 39,228.
Materials and Methods:
Rab11a constructs:
GFP-Rab11a wild-type, constitutively active (Q70L) and dominant negative (S25N) were generously provided by Marino Zerial (Max-Planck
Institute of Molecular Cell Biology and Genetics). mCherry-Rab11a wild-type, constitutively active (S20V) and dominant negative (S25N) were generously provided by Maria Ascaño (Kuruvilla lab, Johns Hopkins University). All plasmids were sequenced prior to use.
Transfections:
Cos7 cells were transfected with the GFP vector, GFP-Rab11a constructs, mCherry vector or the mCherry-Rab11a constructs. 5 x 106 cells in 0.5 ml Opti-
MEM (Gibco) were mixed with 15 µg of the appropriate DNA and electroporated
202 at 240V in an electro cell manipulator 600 (BTX). Transfected cells were diluted into DMEM (Gibco) supplemented with 10% FBS and 1% L-glutamine then grown on cover slips in six-well dishes for 24 hours prior to fixing for
Immunofluorescence staining or the transferrin uptake assay.
Transferrin uptake assay:
To perform the uptake assay, transfected cells were serum starved in
DMEM (Gibco) with 0.2% BSA and 1% L-glutamine for 1 hour at 37°C, then incubated with either Alexa-fluor 546 or 488 conjugated transferrin, 5 mg/ml in
DMEM (Gibco) with 0.2% BSA and 1% L-glutamine for 60 minutes at 37°C. Cells were fixed with 4% formaldehyde in PBS for 15 min at 37°C, at 0, 20 and 60 minutes, and stored at 4°C in PBS for the remainder of the time course. After continuously labeling with transferrin, the media was exchanged to DMEM with
10% FBS and 1% L-glutamine for 60 minutes, after which cells were fixed with
4% formaldehyde in PBS.
The fixed cover slips were incubated in blocking buffer, 0.1% Saponin in
PBS with 1% BSA, for 15 minutes at room temperature to permeabilize the membrane. To label the nuclei, cells were incubated in 1% BSA in PBS with
DAPI (1 µg/ml), then washed with 1% BSA in PBS. Cover slips were mounted using Fluoromount (Sigma) and dried in the dark overnight.
203 27A Immunofluorescence:
Transfected cells were fixed with 4% formaldehyde in PBS for 15 min at
37°C, washed with PBS, then permeabilized with 0.1% Saponin in PBS with 1%
BSA, for 15 minutes at room temperature. Cover slips were placed in a humid chamber and incubated with the 27A mAb in 1% BSA in PBS for 60 minutes, washed three times in 1% BSA in PBS, then incubated with either Alexa-546 or -
488 conjugated goat-anti-mouse secondary antibody for 30 minutes. The cover slips were washed three times in 1% BSA in PBS, with DAPI (1 mg/ml) included in the penultimate wash, then mounted with Fluoromount (Sigma) and dried overnight in the dark.
Immunofluorescence Microscopy:
Immunofluorescence microscopy was performed on an Axiovert 100 LM microscope (Carl Zeiss), equipped with a CoolSnap digital monochrome camera.
Images were processed with Slidebook and Adobe Photoshop. For analysis of the p27 cluster 100-200 cells were examined per condition, with three independent replicates.
204 References:
1 Verhey, K. J. & Hammond, J. W. in Nature Reviews Molecular Cell Biology Vol. 10 765-777 (2009).
2 Torisawa, T. et al. in Nature Publishing Group Vol. 16 1118-1124 (Nature Publishing Group, 2014).
3 Bingham, J. B., King, S. J. & Schroer, T. A. in Meth. Enzymol. Vol. 298 171-184 (1998).
4 Amos, L. A. in J. Cell. Sci. Vol. 93 ( Pt 1) 19-28 (1989).
5 Yeh, E., Skibbens, R. V., Cheng, J. W., Salmon, E. D. & Bloom, K. in J. Cell Biol. Vol. 130 687-700 (1995).
6 Oakley, B. R. & Morris, N. R. in Cell Vol. 19 255-262 (1980).
7 Plamann, M., Minke, P. F., Tinsley, J. H. & Bruno, K. S. in J. Cell Biol. Vol. 127 139-149 (1994).
8 Kon, T. et al. in Nature Vol. 484 345-350 (Nature Publishing Group, 2012).
9 Harte, P. J. & Kankel, D. R. in Genetics Vol. 101 477-501 (1982).
10 Fletcher, D. A. & Mullins, R. D. in Nature Vol. 463 485-492 (2010).
11 Granger, E., McNee, G., Allan, V. & Woodman, P. in Semin. Cell Dev. Biol. Vol. 31 20-29 (2014).
12 Schroer, T. A. & Sheetz, M. P. in J. Cell Biol. Vol. 115 1309-1318 (1991).
13 Carlier, M.-F. et al. in Cell. Mol. Life Sci. Vol. 72 3051-3067 (Springer Basel, 2015).
14 Mandelkow, E. M., Mandelkow, E. & Milligan, R. A. in J. Cell Biol. Vol. 114 977-991 (1991).
15 Vale, R. D. in Cell Vol. 112 467-480 (2003).
16 O'Connell, C. B., Tyska, M. J. & Mooseker, M. S. in Biochim. Biophys. Acta Vol. 1773 615-630 (2007).
17 Krendel, M. & Mooseker, M. S. in Physiology (Bethesda) Vol. 20 239-251 (2005).
205 18 Miki, H., Setou, M., Kaneshiro, K. & Hirokawa, N. in Proceedings of the National Academy of Sciences Vol. 98 7004-7011 (2001).
19 Hirokawa, N. & Noda, Y. in Physiological Reviews Vol. 88 1089-1118 (2008).
20 Hirokawa, N., Noda, Y., Tanaka, Y. & Niwa, S. in Nature Reviews Molecular Cell Biology Vol. 10 682-696 (2009).
21 Gibbons, I. R. in Cell Struct. Funct. Vol. 21 331-342 (1996).
22 Hook, P. & Vallee, R. B. in J. Cell. Sci. Vol. 119 4369-4371 (2006).
23 Schroer, T. A., Steuer, E. R. & Sheetz, M. P. in Cell Vol. 56 937-946 (1989).
24 Paschal, B. M. et al. in Nature Vol. 330 672-674 (1987).
25 Li, M., McGrail, M., Serr, M. & Hays, T. S. in J. Cell Biol. Vol. 126 1475- 1494 (1994).
26 Duncan, J. E. & Warrior, R. in Current Biology Vol. 12 1982-1991 (2002).
27 Swan, A., Nguyen, T. & Suter, B. in Nat. Cell Biol. Vol. 1 444-449 (1999).
28 Galic, M. & Matis, M. in Bioessays Vol. 37 678-686 (2015).
29 Li, Y. Y., Yeh, E., Hays, T. & Bloom, K. in Proceedings of the National Academy of Sciences Vol. 90 10096-10100 (1993).
30 Sullivan, D. S. & Huffaker, T. C. in J. Cell Biol. Vol. 119 379-388 (1992).
31 Beaudouin, J., Gerlich, D., Daigle, N., Eils, R. & Ellenberg, J. in Cell Vol. 108 83-96 (2002).
32 Quintyne, N. J. in J. Cell Biol. Vol. 159 245-254 (2002).
33 Salina, D. et al. in Cell Vol. 108 97-107 (2002).
34 Vaisberg, E. A., Koonce, M. P. & McIntosh, J. R. in J. Cell Biol. Vol. 123 849-858 (1993).
35 Eshel, D. et al. in Proceedings of the National Academy of Sciences Vol. 90 11172-11176 (1993).
36 Yount, A. L., Zong, H. & Walczak, C. E. in Exp. Cell Res. Vol. 334 70-77 (2015).
206 37 Hobdy-Henderson, K. C., Hales, C. M., Lapierre, L. A., Cheney, R. E. & Goldenring, J. R. in Traffic Vol. 4 681-693 (2003).
38 Wilson, G. M. et al. in Mol. Biol. Cell Vol. 16 849-860 (2005).
39 Barr, F. A. & Gruneberg, U. in Cell Vol. 131 847-860 (2007).
40 Fielding, A. B. et al. in EMBO J. Vol. 24 3389-3399 (2005).
41 Vallee, R. B., Wall, J. S., Paschal, B. M. & Shpetner, H. S. in Nature Vol. 332 561-563 (1988).
42 King, S. M., Patel-King, R. S., Wilkerson, C. G. & Witman, G. B. in J. Cell Biol. Vol. 131 399-409 (1995).
43 King, S. M. et al. in Journal of Biological Chemistry Vol. 271 19358- 19366 (1996).
44 King, S. M. et al. in Biochemistry Vol. 37 15033-15041 ( American Chemical Society, 1998).
45 Bowman, A. B. et al. in J. Cell Biol. Vol. 146 165-180 (1999).
46 Benashski, S. E., Harrison, A., Patel-King, R. S. & King, S. M. in Journal of Biological Chemistry Vol. 272 20929-20935 (1997).
47 Takahashi, Y., Edamatsu, M. & Toyoshima, Y. Y. in Proceedings of the National Academy of Sciences Vol. 101 12865-12869 (2004).
48 Kon, T., Nishiura, M., Ohkura, R., Toyoshima, Y. Y. & Sutoh, K. in Biochemistry Vol. 43 11266-11274 (2004).
49 Gee, M. A., Heuser, J. E. & Vallee, R. B. in Nature Vol. 390 636-639 (Nature Publishing Group, 1997).
50 Numata, N. et al. in Biochem. Soc. Trans Vol. 36 131-135 (2008).
51 Kon, T., Mogami, T., Ohkura, R., Nishiura, M. & Sutoh, K. in Nat. Struct. Mol. Biol. Vol. 12 513-519 (2005).
52 Roberts, A. J. et al. in Cell Vol. 136 485-495 (Elsevier Inc., 2009).
53 Carter, A. P., Cho, C., Jin, L. & Vale, R. D. in Science Vol. 331 1159- 1165 (2011).
54 Carter, A. P. et al. in Science Vol. 322 1691-1695 (2008).
55 Gibbons, I. R. et al. in Journal of Biological Chemistry Vol. 280 23960- 23965 (2005).
207 56 Carter, A. P. & Vale, R. D. in Biochem. Cell Biol. Vol. 88 15-21 (2010).
57 Urnavicius, L. et al. in Science Vol. 347 1441-1446 (American Association for the Advancement of Science, 2015).
58 Tynan, S. H., Gee, M. A. & Vallee, R. B. in Journal of Biological Chemistry Vol. 275 32769-32774 (2000).
59 King, S. J., Bonilla, M., Rodgers, M. E. & Schroer, T. A. in Protein Sci. Vol. 11 1239-1250 (Cold Spring Harbor Laboratory Press, 2002).
60 Chowdhury, S., Ketcham, S. A., Schroer, T. A. & Lander, G. C. in Nat. Struct. Mol. Biol. Vol. 22 345-347 (Nature Publishing Group, 2015).
61 Lye, R. J., Porter, M. E., Scholey, J. M. & McIntosh, J. R. in Cell Vol. 51 309-318 (1987).
62 Shpetner, H. S., Paschal, B. M. & Vallee, R. B. in J. Cell Biol. Vol. 107 1001-1009 (1988).
63 Muhua, L., Karpova, T. S. & Cooper, J. A. in Cell Vol. 78 669-679 (1994).
64 Clark, S. W. & Meyer, D. I. in J. Cell Biol. Vol. 127 129-138 (1994).
65 McMillan, J. N. & Tatchell, K. in J. Cell Biol. Vol. 125 143-158 (1994).
66 Kahana, J. A. et al. in Mol. Biol. Cell Vol. 9 1741-1756 (1998).
67 Geiser, J. R. et al. in Mol. Biol. Cell Vol. 8 1035-1050 (1997).
68 Morris, N. R. in Exp. Cell Res. Vol. 98 204-210 (1976).
69 Garnjobst, L. & Tatum, E. L. in Genetics Vol. 57 579-604 (1967).
70 Xiang, X., Beckwith, S. M. & Morris, N. R. in Proceedings of the National Academy of Sciences Vol. 91 2100-2104 (1994).
71 Xiang, X., Osmani, A. H., Osmani, S. A., Xin, M. & Morris, N. R. in Mol. Biol. Cell Vol. 6 297-310 (1995).
72 Bruno, K. S., Tinsley, J. H., Minke, P. F. & Plamann, M. in Proceedings of the National Academy of Sciences Vol. 93 4775-4780 (1996).
73 Beckwith, S. M., Roghi, C. H., Liu, B. & Ronald Morris, N. in J. Cell Biol. Vol. 143 1239-1247 (1998).
74 Xiang, X., Zuo, W., Efimov, V. P. & Morris, N. R. in Curr. Genet. Vol. 35 626-630 (1999).
208 75 Minke, P. F., Lee, I. H. & Plamann, M. in Fungal Genet. Biol. Vol. 28 55- 67 (1999).
76 Efimov, V. P. & Morris, N. R. in J. Cell Biol. Vol. 150 681-688 (2000).
77 Zhang, J., Han, G. & Xiang, X. in Mol. Microbiol. Vol. 44 381-392 (2002).
78 Zhang, J., Li, S., Fischer, R. & Xiang, X. in Mol. Biol. Cell Vol. 14 1479- 1488 (American Society for Cell Biology, 2003).
79 Liu, B., Xiang, X. & Lee, Y.-R. J. in Mol. Microbiol. Vol. 47 291-301 (2003).
80 Zhang, J. et al. in Traffic Vol. 9 1073-1087 (Blackwell Publishing Ltd, 2008).
81 Zhang, J., Li, S., Musa, S., Zhou, H. & Xiang, X. in J. Biol. Chem. Vol. 284 34760-34768 (American Society for Biochemistry and Molecular Biology, 2009).
82 Plough, H. H. & Ives, P. T. in Genetics Vol. 20 42-69 (1935).
83 Swaroop, A., Swaroop, M. & Garen, A. in Proceedings of the National Academy of Sciences Vol. 84 6501-6505 (1987).
84 Gill, S. R. et al. in J. Cell Biol. Vol. 115 1639-1650 (1991).
85 Holzbaur, E. L. et al. in Nature Vol. 360 695-695 (1992).
86 Waterman-Storer, C. M. & Holzbaur, E. L. in Journal of Biological Chemistry Vol. 271 1153-1159 (1996).
87 Eckley, D. M. et al. in J. Cell Biol. Vol. 147 307-320 (1999).
88 Eckley, D. M. & Schroer, T. A. in Mol. Biol. Cell Vol. 14 2645-2654 (2003).
89 Lee, I. H., Kumar, S. & Plamann, M. in Mol. Biol. Cell Vol. 12 2195-2206 (2001).
90 Vierula, P. J. & Mais, J. M. in Mol. Microbiol. Vol. 24 331-340 (1997).
91 Zhang, J. et al. in J. Cell Biol. Vol. 193 1245-1255 (Rockefeller Univ Press, 2011).
92 Sönnichsen, B. et al. in Nature Vol. 434 462-469 (2005).
93 Huang, J., Roberts, A. J., Leschziner, A. E. & Reck-Peterson, S. L. in Cell Vol. 150 975-986 (2012).
209 94 Toropova, K. et al. in Elife Vol. 3 (2014).
95 Derewenda, U. et al. in Structure Vol. 15 1467-1481 (2007).
96 McKenney, R. J., Huynh, W., Tanenbaum, M. E., Bhabha, G. & Vale, R. D. in Science Vol. 345 337-341 (American Association for the Advancement of Science, 2014).
97 Schlager, M. A., Hoang, H. T., Urnavicius, L., Bullock, S. L. & Carter, A. P. in EMBO J. Vol. 33 1855-1868 (2014).
98 Xiang, X., Roghi, C. & Morris, N. R. in Proceedings of the National Academy of Sciences Vol. 92 9890-9894 (1995).
99 Reiner, O. et al. in Nature Vol. 364 717-721 (Nature Publishing Group, 1993).
100 Morris, N. R., Efimov, V. P. & Xiang, X. in Trends Cell Biol. Vol. 8 467- 470 (1998).
101 Wynshaw-Boris, A. in Clin. Genet. Vol. 72 296-304 (Blackwell Publishing Ltd, 2007).
102 Reiner, O. & Sapir, T. in Curr. Opin. Neurobiol. Vol. 23 951-956 (2013).
103 Fry, A. E., Cushion, T. D. & Pilz, D. T. in Am J Med Genet C Semin Med Genet Vol. 166C 198-210 (2014).
104 Lambert de Rouvroit, C. & Goffinet, A. M. in Mech. Dev. Vol. 105 47-56 (2001).
105 Sheeman, B. et al. in Current Biology Vol. 13 364-372 (2003).
106 Hebbar, S. et al. in J. Cell Biol. Vol. 182 1063-1071 (Rockefeller Univ Press, 2008).
107 Yingling, J. et al. in Cell Vol. 132 474-486 (2008).
108 Zimdahl, B. et al. in Nat Genet Vol. 46 245-252 (2014).
109 Wang, S. & Zheng, Y. in Journal of Biological Chemistry Vol. 286 587- 593 (2010).
110 Wang, S. et al. in Mol. Biol. Cell Vol. 24 3522-3533 (American Society for Cell Biology, 2013).
111 McKenney, R. J., Vershinin, M., Kunwar, A., Vallee, R. B. & Gross, S. P. in Cell Vol. 141 304-314 (Elsevier Ltd, 2010).
210 112 Schafer, D. A., Gill, S. R., Cooper, J. A., Heuser, J. E. & Schroer, T. A. in J. Cell Biol. Vol. 126 403-412 (Rockefeller Univ Press, 1994).
113 King, S. J., Brown, C. L., Maier, K. C., Quintyne, N. J. & Schroer, T. A. in Mol. Biol. Cell Vol. 14 5089-5097 (2003).
114 Siglin, A. E. et al. in PLoS ONE Vol. 8 e59453 (Public Library of Science, 2013).
115 Culver Hanlon, T. L., Lex, S. A., Stephens, A. D., Quintyne, N. J. & King, S. J. in Nat. Cell Biol. Vol. 8 264-270 (2006).
116 Waterman-Storer, C. M., Karki, S. & Holzbaur, E. L. in Proceedings of the National Academy of Sciences Vol. 92 1634-1638 (1995).
117 Vaughan, K. T. & Vallee, R. B. in J. Cell Biol. Vol. 131 1507-1516 (1995).
118 Karki, S. & Holzbaur, E. L. in Journal of Biological Chemistry Vol. 270 28806-28811 (1995).
119 Moughamian, A. J. & Holzbaur, E. L. F. in Neuron Vol. 74 331-343 (Elsevier Inc., 2012).
120 Moughamian, A. J., Osborn, G. E., Lazarus, J. E., Maday, S. & Holzbaur, E. L. F. in J. Neurosci. Vol. 33 13190-13203 (Society for Neuroscience, 2013).
121 Cheong, F. K. Y., Feng, L., Sarkeshik, A., Yates, J. R. & Schroer, T. A. in Mol. Biol. Cell Vol. 25 2171-2180 (American Society for Cell Biology, 2014).
122 Imai, H., Narita, A., Maéda, Y. & Schroer, T. A. in J. Mol. Biol. Vol. 426 3262-3271 (2014).
123 Hoogenraad, C. C. et al. in EMBO J. Vol. 20 4041-4054 (2001).
124 Matanis, T. et al. in Nat. Cell Biol. Vol. 4 986-992 (2002).
125 Splinter, D. et al. in Mol. Biol. Cell Vol. 23 4226-4241 (2012).
126 Schroer, T. A. & Sheetz, M. P. in Annu. Rev. Physiol. Vol. 53 629-652 (1991).
127 Blocker, A. et al. in J. Cell Biol. Vol. 137 113-129 (1997).
128 Presley, J. F. et al. in Nature Vol. 389 81-85 (1997).
129 Xiang, X. et al. in Cell. Mol. Life Sci. 1-14 (Springer Basel, 2015).
211 130 Vallee, R. B., McKenney, R. J. & Ori-McKenney, K. M. in Nature Publishing Group Vol. 14 224-230 (2012).
131 Gaglio, T. et al. in J. Cell Biol. Vol. 135 399-414 (1996).
132 Wordeman, L., Steuer, E. R., Sheetz, M. P. & Mitchison, T. in J. Cell Biol. Vol. 114 285-294 (1991).
133 Steuer, E. R., Wordeman, L., Schroer, T. A. & Sheetz, M. P. in Nature Vol. 345 266-268 (1990).
134 Schroer, T. A. in J. Cell Biol. Vol. 127 1-4 (1994).
135 Pfarr, C. M. et al. in Nature Vol. 345 263-265 (1990).
136 Osmani, A. H., Osmani, S. A. & Morris, N. R. in J. Cell Biol. Vol. 111 543- 551 (1990).
137 Koonce, M. P., Grissom, P. M. & McIntosh, J. R. in J. Cell Biol. Vol. 119 1597-1604 (1992).
138 Hyman, A. A. & White, J. G. in J. Cell Biol. Vol. 105 2123-2135 (1987).
139 Hyman, A. A., Middleton, K., Centola, M., Mitchison, T. J. & Carbon, J. in Nature Vol. 359 533-536 (1992).
140 Hyman, A. A. in J. Cell Biol. Vol. 109 1185-1193 (1989).
141 Fyrberg, C., Ryan, L., Kenton, M. & Fyrberg, E. in J. Mol. Biol. Vol. 241 498-503 (1994).
142 Fu, C. et al. in Eukaryotic Cell Vol. 10 1100-1109 (American Society for Microbiology, 2011).
143 Farkasovsky, M. & Küntzel, H. in J. Cell Biol. Vol. 131 1003-1014 (1995).
144 Egan, M. J., McClintock, M. A. & Reck-Peterson, S. L. in Curr. Opin. Microbiol. Vol. 15 637-645 (2012).
145 Li, J., Lee, W.-L. & Cooper, J. A. in Nat. Cell Biol. Vol. 7 686-690 (Nature Publishing Group, 2005).
146 Métin, C., Vallee, R. B., Rakic, P. & Bhide, P. G. in J. Neurosci. Vol. 28 11746-11752 (Society for Neuroscience, 2008).
147 Tsai, J.-W., Bremner, K. H. & Vallee, R. B. in Nat Neurosci Vol. 10 970- 979 (2007).
212 148 Minke, P. F., Lee, I. H., Tinsley, J. H., Bruno, K. S. & Plamann, M. in Mol. Microbiol. Vol. 32 1065-1076 (1999).
149 Cheng, J., Tiyaboonchai, A., Yamashita, Y. M. & Hunt, A. J. in Development Vol. 138 831-837 (The Company of Biologists Limited, 2011).
150 Elias, S. et al. in Stem Cell Reports Vol. 2 491-506 (2014).
151 Molina-Calavita, M. et al. in J. Neurosci. Vol. 34 10034-10040 (Society for Neuroscience, 2014).
152 Sasaki, S. et al. in Neuron Vol. 28 681-696 (2000).
153 Smith, D. S. et al. in Nat. Cell Biol. Vol. 2 767-775 (2000).
154 Zylkiewicz, E. et al. in J. Cell Biol. Vol. 192 433-445 (2011).
155 McKenney, R. J., Weil, S. J., Scherer, J. & Vallee, R. B. in Journal of Biological Chemistry Vol. 286 39615-39622 (2011).
156 Barbar, E. in Biochem. Soc. Trans Vol. 40 1009-1013 (2012).
157 Nyarko, A., Song, Y. & Barbar, E. in J. Biol. Chem. Vol. 287 24884- 24893 (American Society for Biochemistry and Molecular Biology, 2012).
158 Ma, L. et al. in Nature Publishing Group Vol. 11 247-256 (2009).
159 Bloom, G. S., Schoenfeld, T. A. & Vallee, R. B. in J. Cell Biol. Vol. 98 320-330 (1984).
160 Imai, H., Narita, A., Schroer, T. A. & Maéda, Y. in J. Mol. Biol. Vol. 359 833-839 (2006).
161 Steffen, W., Hodgkinson, J. L. & Wiche, G. in Journal of Structural Biology Vol. 117 227-235 (1996).
162 Hodgkinson, J. L., Peters, C., Kuznetsov, S. A. & Steffen, W. in Proc. Natl. Acad. Sci. U.S.A. Vol. 102 3667-3672 (National Acad Sciences, 2005).
163 Yeh, T.-Y., Quintyne, N. J., Scipioni, B. R., Eckley, D. M. & Schroer, T. A. in Mol. Biol. Cell Vol. 23 3827-3837 (American Society for Cell Biology, 2012).
164 Burkhardt, J. K., Echeverri, C. J., Nilsson, T. & Vallee, R. B. in J. Cell Biol. Vol. 139 469-484 (1997).
165 Quintyne, N. J. et al. in J. Cell Biol. Vol. 147 321-334 (1999).
213 166 Valetti, C. et al. in Mol. Biol. Cell Vol. 10 4107-4120 (1999).
167 King, S. J. & Schroer, T. A. in Nat. Cell Biol. Vol. 2 20-24 (2000).
168 Tribet, C., Audebert, R. & Popot, J. L. in Proceedings of the National Academy of Sciences Vol. 93 15047-15050 (1996).
169 Flötenmeyer, M., Weiss, H., Tribet, C., Popot, J.-L. & Leonard, K. in J Microsc Vol. 227 229-235 (Blackwell Publishing Ltd, 2007).
170 Yamashita, A., Maeda, K. & Maéda, Y. in EMBO J. Vol. 22 1529-1538 (2003).
171 Narita, A., Takeda, S., Yamashita, A. & Maéda, Y. in EMBO J. Vol. 25 5626-5633 (2006).
172 Yeh, T.-Y. et al. in EMBO J. Vol. 32 1023-1035 (2013).
173 Lees-Miller, J. P., Helfman, D. M. & Schroer, T. A. in Nature Vol. 359 244-246 (Nature Publishing Group, 1992).
174 Karki, S., LaMonte, B. & Holzbaur, E. L. in J. Cell Biol. Vol. 142 1023- 1034 (1998).
175 Echeverri, C. J., Paschal, B. M., Vaughan, K. T. & Vallee, R. B. in J. Cell Biol. Vol. 132 617-633 (1996).
176 Paschal, B. M. et al. in Journal of Biological Chemistry Vol. 268 15318- 15323 (1993).
177 Melkonian, K. A., Maier, K. C., Godfrey, J. E., Rodgers, M. & Schroer, T. A. in Journal of Biological Chemistry Vol. 282 19355-19364 (2007).
178 Maier, K. C., Godfrey, J. E., Echeverri, C. J., Cheong, F. K. Y. & Schroer, T. A. in Traffic Vol. 9 481-491 (2008).
179 Amaro, I. A., Costanzo, M., Boone, C. & Huffaker, T. C. in Genetics Vol. 178 703-709 (2008).
180 Bhabha, G., Johnson, G. T., Schroeder, C. M. & Vale, R. D. in Trends Biochem. Sci. Vol. 41 94-105 (Elsevier, 2016).
181 Carter, A. P., Diamant, A. G. & Urnavicius, L. in Current Opinion in Structural Biology Vol. 37 62-70 (2016).
182 Cianfrocco, M. A., DeSantis, M. E., Leschziner, A. E. & Reck-Peterson, S. L. in Annu. Rev. Cell Dev. Biol. Vol. 31 83-108 ( Annual Reviews, 2015).
183 Roberts, A. J. et al. in Structure Vol. 20 1670-1680 (Elsevier, 2012).
214 184 Li, X.-J. & Li, S.-H. in Trends Pharmacol. Sci. Vol. 26 1-3 (Elsevier, 2005).
185 Lin, X. et al. in Sci Rep Vol. 4 7282 (2014).
186 Loss, O. & Stephenson, F. A. in J. Neurosci. Res. Vol. 93 1056-1066 (2015).
187 Matsui, T., Ohbayashi, N. & Fukuda, M. in J. Biol. Chem. Vol. 287 28619- 28631 (American Society for Biochemistry and Molecular Biology, 2012).
188 Ogawa, F. et al. in Human Molecular Genetics Vol. 23 906-919 (Oxford University Press, 2014).
189 Yadav, S., Puthenveedu, M. A. & Linstedt, A. D. in Developmental Cell Vol. 23 153-165 (Elsevier, 2012).
190 Egan, M. J., Tan, K. & Reck-Peterson, S. L. in J. Cell Biol. Vol. 197 971- 982 (Rockefeller Univ Press, 2012).
191 Hayashi, I., Plevin, M. J. & Ikura, M. in Nat. Struct. Mol. Biol. Vol. 14 980- 981 (2007).
192 Spector, I., Shochet, N. R., Blasberger, D. & Kashman, Y. in Cell Motil. Cytoskeleton Vol. 13 127-144 (Wiley Subscription Services, Inc., A Wiley Company, 1989).
193 Spector, I., Shochet, N. R., Kashman, Y. & Groweiss, A. in Science Vol. 219 493-495 (1983).
194 Carter, S. B. in Nature Vol. 213 261-264 (1967).
195 Brenner, S. L. & Korn, E. D. in Journal of Biological Chemistry Vol. 255 841-844 (1980).
196 MacLean-Fletcher, S. & Pollard, T. D. in Cell Vol. 20 329-341 (1980).
197 Selden, L. A., Gershman, L. C. & Estes, J. E. in Biochemical and Biophysical Research Communications Vol. 95 1854-1860 (1980).
198 Kirschner, M. W., Williams, R. C., Weingarten, M. & Gerhart, J. C. in Proceedings of the National Academy of Sciences Vol. 71 1159-1163 (1974).
199 Wani, M. C., Taylor, H. L., Wall, M. E., Coggon, P. & McPhail, A. T. in J. Am. Chem. Soc. Vol. 93 2325-2327 (1971).
200 Kumar, N. in Journal of Biological Chemistry Vol. 256 10435-10441 (1981).
215 201 Allingham, J. S., Zampella, A., D'Auria, M. V. & Rayment, I. in Proc. Natl. Acad. Sci. U.S.A. Vol. 102 14527-14532 (National Acad Sciences, 2005).
202 Klenchin, V. A. et al. in Nat. Struct. Biol. Vol. 10 1058-1063 (2003).
203 Saito, S. & Karaki, H. in Clin. Exp. Pharmacol. Physiol. Vol. 23 743-746 (1996).
204 Saito, S., Watabe, S., Ozaki, H., Fusetani, N. & Karaki, H. in J. Biol. Chem. Vol. 269 29710-29714 (1994).
205 Hirata, K. et al. in J. Mol. Biol. Vol. 356 945-954 (2006).
206 Wada, S., Matsunaga, S., Saito, S., Fusetani, N. & Watabe, S. in J. Biochem. Vol. 123 946-952 (1998).
207 Hori, M. et al. in FEBS Lett. Vol. 322 151-154 (1993).
208 Okada, T., Narai, A., Matsunaga, S., Fusetani, N. & Shimizu, M. in Toxicol In Vitro Vol. 14 219-226 (2000).
209 Takakuwa, R. et al. in Exp. Cell Res. Vol. 257 238-244 (2000).
210 Lázaro-Diéguez, F. et al. in Cell Motil. Cytoskeleton Vol. 63 778-791 (Wiley Subscription Services, Inc., A Wiley Company, 2006).
211 Castorena-Gonzalez, J. A., Staiculescu, M. C., Foote, C. A., Polo-Parada, L. & Martinez-Lemus, L. A. in Am. J. Physiol. Heart Circ. Physiol. Vol. 306 H485-495 (2014).
212 Nakamura, M. et al. in Acta Med. Okayama Vol. 56 43-50 (2002).
213 Cavolo, S. L. et al. in Mol. Biol. Cell mbc.E14-11-1564 (American Society for Cell Biology, 2015).
214 Martin, M. et al. in Mol. Biol. Cell Vol. 10 3717-3728 (1999).
215 Rogers, S. L., Tint, I. S., Fanapour, P. C. & Gelfand, V. I. in Proceedings of the National Academy of Sciences Vol. 94 3720-3725 (1997).
216 Haghnia, M. et al. in Mol. Biol. Cell Vol. 18 2081-2089 (2007).
217 Brady, S. T., Pfister, K. K. & Bloom, G. S. in Proceedings of the National Academy of Sciences Vol. 87 1061-1065 (1990).
218 Muller, J. et al. in Mol. Biol. Cell Vol. 16 5736-5748 (2005).
216 219 Ozaki, Y. et al. in Journal of Biological Chemistry Vol. 286 5589-5598 (2011).
220 Chen, T.-Y. et al. in J. Cell. Biochem. n/a-n/a (2015).
221 Zerial, M. & McBride, H. in Nature Reviews Molecular Cell Biology Vol. 2 107-117 (Nature Publishing Group, 2001).
222 Green, E. G. et al. in Biochemical and Biophysical Research Communications Vol. 239 612-616 (1997).
223 Hopkins, C. R., Gibson, A., Shipman, M., Strickland, D. K. & Trowbridge, I. S. in J. Cell Biol. Vol. 125 1265-1274 (1994).
224 Ullrich, O., Reinsch, S., Urbé, S., Zerial, M. & Parton, R. G. in J. Cell Biol. Vol. 135 913-924 (1996).
225 Wang, X. in Journal of Biological Chemistry Vol. 275 29138-29146 (2000).
226 Horgan, C. P. et al. in Traffic Vol. 8 414-430 (2007).
227 Hales, C. M. et al. in Journal of Biological Chemistry Vol. 276 39067- 39075 (2001).
228 Montagnac, G., Echard, A. & Chavrier, P. in Current Opinion in Cell Biology Vol. 20 454-461 (2008).
217
Stephanie A. Ketcham
Department of Biology The Johns Hopkins University 3400 N. Charles Street, Mudd Hall room 209 Baltimore MD, 21218 (410)516-5374 Email: [email protected]
EDUCATION: 2008-present The Johns Hopkins University, Baltimore, MD Ph.D. program in Cell, Molecular, Developmental Biology and Biophysics
2004-2008 Saint Michael’s College, Colchester, VT B.S., Cellular and Molecular Biology Graduated Magna cum laude
Research Experience 2009-present Doctoral research with Dr. Trina Schroer Department of Biology, The Johns Hopkins University Structural and biochemical analysis of the dynein motor.
2008-2009 Rotations with: Dr. David Zapulla: examining Telomerase function in S. cerevisiae. Dr. Trina Schroer: Dynactin p27 functions in cultured cells. Dr. Karen Beemon: looking for small RNAs in P. falciparum. Dr. Beverly Wendland: characterizing protein interactions necessary for endocytosis in S. cerevisiae.
2007-2008 Undergraduate research with Dr. Mark Lubkowitz Department of Biology, Saint Michael’s College Characterization of oligopeptide transport in rice.
2006-2008 Internship at Green Mountain Antibodies, Burlington, VT Monoclonal antibody production for commercial and research purposes.
HONORS AND AWARDS: 2007 Member, Delta Epsilon Sigma 2008 Member, Phi Beta Kappa 2008 Member, Sigma Xi 2013 Graduate Travel Award (American Society for Cell Biology)
218 PUBLICATIONS:
Wang, S. Ketcham, S.A. Schön, A. Goodman, B. Wang, Y. Yates, J. 3rd, Freire, E. Schroer, T.A. Zheng, Y. (2013) Nudel/NudE and Lis1 promote dynein and dynactin interactions in the context of spindle morphogenesis. Mol. Biol. Cell.
Chowdhury, S. Ketcham, S.A. Schroer, T.A. Lander, G.C. (2015) Structural organization of the dynein-dynactin complex bout to microtubules. Nat. Struct. Mol. Biol.
Cavolo, S. Zhou, C. Ketcham, S.A. Suzuki, M. Kresimir, U. Silverman, M. Schroer, T.A. Levitan, E. (2015) Mycalolide B dissociates dynactin and abolishes retrograde axonal transport of dense-core vesicles. Mol. Biol. Cell.
Susanne Höing, S. Yeh, T.Y. Baumann, M. Martinez, N. Habenberger, P. Kremer, L. Drexler, H. Küchler, P. Reinhardt, P. Choidas, A. Zischinsky, M.L. Zischinsky, G. Ketcham, S.A. Wagner, L. Nussbaumer, P. Klebl, B. Ziegler, S. Schroer, T.A. Schöler, H. Waldmann, H. Dynarrestin, a Novel Dynein Inhibitor that does not Block Ciliogenesis. In review, Nature Chem. Biol.
Ketcham, S.A. Schroer, T.A. Characterization of the protein-protein interactions within the dynactin shoulder. Manuscript in preparation.
POSTER PRESENTATIONS:
Protein-protein interactions within dynactin’s shoulder/sidearm domain. Stephanie A. Ketcham and Trina A. Schroer 50th American Society for Cell Biology Annual Meeting, Philadelphia, PA, 2010
Protein-protein interactions within dynactin’s shoulder/sidearm. Stephanie A. Ketcham and Trina A. Schroer 51st American Society for Cell Biology Annual Meeting, Denver, CO, 2011
Probing the subunit interactions in the dynactin shoulder/sidearm. Stephanie A. Ketcham and Trina A. Schroer 52nd American Society for Cell Biology Annual Meeting, San Francisco, CA 2012
Interactions between cytoplasmic dynein and the regulatory complexes: dynactin and Lis1-Nudel. Stephanie A. Ketcham and Trina A. Schroer 53rd American Society for Cell Biology Annual Meeting, New Orleans, LA, 2013
Mapping of the dynactin shoulder domain aimed at understanding its three-helix bundle structure Stephanie A. Ketcham and Trina A. Schroer 55th American Society for Cell Biology Annual Meeting, San Diego, CA 2015
219 TEACHING EXPERIENCE:
Teaching Assistant, Biochemistry laboratory, The Johns Hopkins University, Fall 2009. Lectured on and demonstrated laboratory techniques and assisted students in weekly laboratory experiments. Graded weekly laboratory reports, weekly quizzes and exams. Dr. Robert Horner, Department of Biology.
Teaching Assistant, Cell Biology, The Johns Hopkins University, Spring 2010. Lectured on and demonstrated laboratory techniques and assisted students in weekly laboratory experiments. Graded weekly laboratory reports, weekly quizzes and exams. Dr. Robert Horner, Department of Biology.
Teaching Assistant, Cell Biology, The Johns Hopkins University, Spring 2012, 2014 and 2015. Helped review exams for clarity of questions prior to administration and graded exams for a class of 100-200 students. Dr. Kathryn Tifft Oshinnaiye, Dr. Emily Fisher, Dr. Trina Schroer, Dr. Kyle Cunningham, Department of Biology.
Teaching Assistant, Biochemistry, The Johns Hopkins University, Fall 2012, 2013 -2015. Helped review exams for clarity of questions prior to administration and graded exams for a class of 100-200 students. Dr. Kathryn Tifft Oshinnaiye, Dr. Emily Fisher, Dr. Vince Hilser, Dr. Young Sam Lee, Dr. Christian Kaiser, Department of Biology.
Teaching Assistant, Immunology, The Johns Hopkins University, Spring 2013. Held weekly office hours, graded homework and exams and offered one-on-one feedback on exams and homework for a class of 25 students. Dr. Michael Edidin, Department of Biology.
Teaching Assistant, AIDS, The Johns Hopkins University, Spring 2016. Held weekly office hours, graded homework and exams and offered one-on-one feedback on exams and homework for a class of 50 students. Dr. Trina A Schroer, Department of Biology
LETTERS (BY REQUEST):
Trina A. Schroer: Department of Biology, Johns Hopkins University ([email protected])
Yixian Zheng: Carnegie Institute of Embryology ([email protected])
Gabriel C. Lander: Integrative Structural and Computational Biology, The Scripps Research Institute ([email protected]
220