Mechanisms of Axonal Transport in ALS

Aus der Klinik und Poliklinik für Neurologie

Direktor: Herr Prof. Dr. Heinz Reichmann

Mechanisms of Axonal Transport Defects in ALS

Dissertationsschrift

Zur Erlangung des akademischen Grades

Doktor der Biomedizin

Doctor rerum medicinalium (Dr. rer. medic.)

vorgelegt bei

der Medizinischen Fakultät Carl Gustav Carus

der Technischen Universität Dresden

von

M.Sc. Anne Seifert

geboren am 05.07.1992 in Dresden

Dresden, Januar 2021

Für meine Mama.

i Anne Seifert

Gutachter:

1. Gutachter: Prof. Dr. Dr. Andreas Hermann, Sektion für Translationale Neurodegeneration

"Albrecht Kossel", Klinik und Poliklinik für Neurologie, Universitätsmedizin

Rostock, Gehlsheimer Straße 20, 18147 Rostock, Germany

2. Gutachter: Prof. Dr. Stefan Diez, Technische Universität Dresden, Center for Molecular and

Cellular Bioengineering (CMCB), B CUBE - Center for Molecular Bioengineering,

Tatzberg 41, 01307 Dresden, Germany

Weitere Mitglieder der Prüfungskommission:

Prof. Dr. Gerd Kempermann (Vorsitzender)

Prof. Dr. Björn Falkenburger

Prof. Dr. Mike O. Karl

Datum der Verteidigung: 23.04.2021

The work described in this thesis was performed at the Universitätsklinikum Carl Gustav Carus,

Fetscherstraße 74, 01307 Dresden and the B CUBE - Center for Molecular Bioengineering,

Technische Universität Dresden, Tatzberg 41, 01307 Dresden.

Mechanisms of Axonal Transport in ALS Table of Contents

Anne Seifert ii

Table of Contents List of Abbreviations ................................................................................................................... iv

1. Introduction ............................................................................................................................ 1

1.1 Amyotrophic lateral sclerosis (ALS) .................................................................................. 1

1.2 Axonal transport ............................................................................................................... 9

1.3 Modelling ALS and axonal transport in vitro .....................................................................18

1.4 Aim of this study ..............................................................................................................22

2. Materials and Methods ..........................................................................................................24

2.1 Materials ..........................................................................................................................24

2.2 Methods ..........................................................................................................................32

3. Results ..................................................................................................................................46

3.1 Modification of a kinesin-1-dependent microtubule gliding assay for the application of

whole cell lysates ..................................................................................................................46

3.2 Determination of assay sensitivity with the microtubule-associated protein tau ................53

3.3 Investigation of a direct interference of recombinant FUS-GFP with kinesin-1 motility and

microtubule gliding.................................................................................................................54

3.4 Kinesin-1-dependent microtubule gliding and assay sensitivity in the presence of whole

cell lysates expressing GFP-labelled FUS variants ................................................................58

3.5 Arrest in gliding of single microtubules caused by Tau-GFP compared to aging of the

assay .....................................................................................................................................60

3.6 Differential effects of 2N3R and 2N4R tau individually on kinesin-1-dependent microtubule

gliding velocity and microtubule binding .................................................................................64

3.7 Impact of increasing 4R:3R tau isoform ratios on kinesin-1-dependent microtubule gliding

and its microtubule binding ....................................................................................................68

3.8 Expression levels of 4R and 3R tau isoforms in whole cell lysates ..................................71

4. Discussion ............................................................................................................................73

4.1 The modified kinesin-1-dependent microtubule gliding assay detects nanomolar amounts

of recombinant human tau-GFP.............................................................................................74

4.2 Wildtype and ALS-associated FUS variants do not directly interfere with kinesin-1 motility

and do not bind to microtubules .............................................................................................76

4.3 An increase in 4R:3R tau isoform ratio might contribute to the axonal transport defects

observed in FUS-ALS ............................................................................................................79

4.4 FUS variants may indirectly affect microtubule-based axonal transport by acting on a

multitude of cellular processes ..............................................................................................88

4.5 Outlook ............................................................................................................................93

4.6 Conclusion .......................................................................................................................94

Mechanisms of Axonal Transport in ALS Table of Contents

Anne Seifert iii

References ...............................................................................................................................95

Supplementary Figures ........................................................................................................... 133

Supplementary Tables ............................................................................................................ 134

List of Figures ......................................................................................................................... 139

List of Tables .......................................................................................................................... 140

List of Supplementary Figures ................................................................................................. 141

List of Supplementary Tables .................................................................................................. 141

Summary ................................................................................................................................ 142

Zusammenfassung ................................................................................................................. 144

Acknowledgements ................................................................................................................. 147

Anlage 1.................................................................................................................................. 149

Anlage 2.................................................................................................................................. 150

Mechanisms of Axonal Transport in ALS List of Abbreviations

Anne Seifert iv

List of Abbreviations

AA ........................................................................................................................... Ascorbig acid ALS .................................................................................................. Amyotrophic lateral sclerosis APC .................................................................................................. Adenomatous polyposis coli APP ..................................................................................................... Amyloid precursor protein BCA ................................................................................................................. Bicinchoninic acid BDNF ........................................................................................ Brain-derived neurotrophic factor C9orf72 ............................................................................ Chromosome 9 open reading frame 72 CNS ........................................................................................................ Central nervous system CV ...................................................................................................................... Column volumes DAPT .................................. N-[N-(3,5-Difluorphenacetyl)-L-alanyl]-S-phenylglycin-tert-butylester DDB ...................................................................................................... Dynein dynactin BICD2N DDS .......................................................................................................... Dichlorodimethylsilane DHCs ........................................................................................................... Dynein heavy chains ER .......................................................................................................... Endoplasmatic reticulum FRET ............................................................................ Fluorescence resonance energy transfer FTD ....................................................................................................... Frontotemporal dementia FUS ................................................................................................................. Fused in sarcoma GMPCPP .................................................................. Guanosine 50-[a,b-methylene] triphosphate HDAC1 ...................................................................................................... Histone deacetylase 1 HDAC6 ...................................................................................................... Histone deacetylase 6 hESC ................................................................................................ Human embryonic stem cell HR .....................................................................................................Homologous recombination HRP ........................................................................................................ Horseradish peroxidase HSP60 ............................................................................................. heat shock protein of 60 kDa iPSCs ............................................................................................. Induced pluripotent stem cells JIP ............................................................................. c-Jun N-terminal kinase-interacting protein KHCs .......................................................................................................... Kinesin heavy chains KLCs .............................................................................................................. Kinesin light chains MAPs ........................................................................................... Microtubule-associated protein Miro .................................................................................................... Mitochondrial Rho GTPase N ................................................................................................................ Amino-terminal insert NDs .................................................................................................. Neurodegenerative diseases NHEJ ................................................................................................ Nonhomologous end joining NMR ................................................................................................ Nuclear magnetic resonance NPCs ..................................................................................................... Neuronal precursor cells PAR ....................................................................................... Poly adenosine diphosphate ribose PARP ................................................................. Poly adenosine diphosphate ribose polymerase PBS ....................................................................................................... Phosphate Buffer Saline PFS ............................................................................................................ Perfect Focus System PLO .................................................................................................................... Poly-L-Ornithine R ........................................................................................................ Microtubule-binding repeat RBPs .......................................................................................................... RNA-binding proteins rhBDNF ..................................................................................... Brain-derived neurotrophic factor rhGDNF ..................................................... Recombinant human glial derived neurotrophic factor RNP ................................................................................................................. Ribonucleoprotein ROS ...................................................................................................... Reactive oxygen species RT ................................................................................................................... Room temperature SAG ............................................................................................................. Smoothened agonist

Mechanisms of Axonal Transport in ALS List of Abbreviations

Anne Seifert v

SDS-PAGE ....................................... Sodium dodecyl sulfate-polyacrylamide gel electrophoresis SEC ............................................................................................ Size exclusion chromatography SEM .................................................................................................... standard error of the mean SFPQ ........................................................................... Splicing factor proline- and glutamine-rich SOD1 ...................................................................................................... Superoxide dismutase 1 TDP-43 ........................................................................................... TAR DNA-Binding Protein 43 TGFβ-3 ........................................................................................ Transforming growth factor β-3 TIRF ..................................................................................... Total internal reflection fluorescence TRAK ................................................................................................................ Trafficking kinase

Mechanisms of Axonal Transport in ALS Introduction

Anne Seifert 1 / 157

1. Introduction

1.1 Amyotrophic lateral sclerosis (ALS)

Neurodegenerative diseases (NDs), such as Alzheimer’s, Parkinson’s, Huntington’s disease,

Frontotemporal dementia (FTD), and ALS, are among the most common causes for mortality

worldwide, with increasing tendency. NDs are highly age-dependent, occurring typically but not

exclusively in the elderly. It becomes progressively important to understand the underlying

pathomechanisms of these diseases, since the elderly population has increased over the last

decades (Heemels, 2016). To date, NDs are generally incurable and only limited treatment

options exist for most of them, partly because the underlying pathophysiology is very diverse.

Some disorders primarily cause cognitive impairment and/or memory loss as seen in FTD and

Alzheimer’s disease (Abeliovich and Gitler, 2016; Canter et al., 2016; Wyss-Coray, 2016), while

others mainly affect the motor system, causing movement, speech, and breathing deficits, as

seen in ALS (Taylor et al., 2016).

ALS is the most common ND specifically affecting cortical (upper) motor neurons in the

primary motor cortex, and spinal (lower) motor neurons in the brainstem and spinal cord. This

disease was first described by Jean-Martin Charcot and Alix Joffroy in 1869 (Charcot and

Joffroy, 1869). Various studies report an incidence for ALS ranging from 0.6 to 3.8 per 100 000

persons/year and a prevalence of between 4.1 and 8.4 per 100 000 persons, with regional

differences (reviewed by Longinetti and Fang, 2019). Men are at higher risk to develop ALS

than women, with reported male-to-female ratios between 1.2 and 1.5 (Manjaly et al., 2010). It is

a progressive disorder with a median age of onset between 51 and 66 years of age throughout

the world (Longinetti and Fang, 2019), in Germany around 61 years (Dorst et al., 2019), and

usually leads to death due to respiratory failure about 2-5 years after symptom onset (Naumann

et al., 2018). ALS commonly manifests as a spinal onset (e.g. weakness in the limbs, up to

82 % of cases), but patients may also show bulbar onset (e.g. difficulty in speaking or

swallowing), mixed spinal and bulbar onset, or other forms of onset, including thoracic onset

(D’Ovidio et al., 2019), dementia or respiratory symptoms (Longinetti et al., 2018; Leighton et

al., 2019). While all ALS patients suffer from motoric symptoms, only 30 % of patients also

develop symptoms of FTD (Lomen-Hoerth, 2011), which also include changes in mood and

behavior (Erkkinen et al., 2018), and those patients were found to face a shorter life expectancy

(Olney et al., 2005).

There is no cure for ALS to date and only two approved drugs, Riluzole and Edaravone.

For a long time, Riluzole, a presumed glutamate antagonist, has been the only authorized drug



for the treatment of ALS after its approval in 1995 (Wokke, 1996). It acts anti-excitotoxic, which

may be related to its ability to inhibit glutamate release, to inactivate voltage-gated sodium

channels, or to interfere with intracellular signal transmission following transmitter binding at

excitatory receptors (Jaiswal, 2019). Recently, Edaravone has been approved in some countries

as an alternative or additional treatment (Bhandari et al., 2018). Edaravone is thought to be a

scavenger for reactive oxygen species, and reportedly eliminates lipid peroxides and hydroxyl

radicals. In neurons, it presumably alleviates injuries caused by free oxygen radicals (Jaiswal,

2019). The exact cellular and molecular targets of Edaravone are however still unknown. Both

Riluzole and Edaravone are effective only in subpopulations of ALS patients and increase

survival time or slow down progression of the disease (Bensimon et al., 1994; Abe et al., 2017;

Jaiswal, 2019).

On the cellular level, ALS is a non-cell-autonomous disease and characterized by the

progressive atrophy of cortical and spinal motor neurons, a reduction in size of the remaining

neurons, hyperexcitability of the motor system (Ferraiuolo et al., 2011), as well as astro- and

microgliosis (Saberi et al., 2015). The latter implicates an inflammatory component in ALS,

which has been subject of a variety of studies in the past. ALS-associated mutations in

superoxide dismutase 1 (SOD1) expressed in astrocytes have been linked to enhanced

activation of microglia in a mouse model of ALS, which may play a role in disease progression

due to the enhanced production of nitric oxide and toxic cytokines (Yamanaka et al., 2008). In

contrast, reactive astrocytes and microglia were found to surround degenerating motor neurons

in ALS patients and mouse models (McGeer and McGeer, 2002; Boillee et al., 2006), where

they secreted proinflammatory cytokines (Saberi et al., 2015). This suggests that a

neuroinflammatory response may exert neuroprotective as well as neurodegenerative effects,

and provides additional evidence for an involvement of not only neurons, but also astrocytes,

microglia, and oligodendrocytes in ALS pathology (Scekic-Zahirovic et al., 2017).

On the molecular level, there is a general consensus that multiple pathogenic processes

contribute to the neuropathology of ALS, including abnormal RNA processing (Donnelly et al.,

2014), aberrant protein folding and the formation of stress granules (dense aggregates

containing proteins and RNA forming under stress conditions within the cell, Gutierrez-Beltran et

al., 2015; Parakh & Atkin, 2016), mitochondrial dysfunction (Smith et al., 2019) and impaired

axonal transport (Ferraiuolo et al., 2011; Ikenaka et al., 2012; Sama et al., 2014; De Vos and

Hafezparast, 2017a). The exact type and degree of neuropathology essentially depends on the

underlying cause for ALS.



About 90 % of cases are sporadic, where no specific genetic or environmental cause can

be identified. Common risk factors for sporadic ALS include older age (largely increasing above

age 50) and male sex. Other factors, such as physical fitness, repeated head injury,

occupational and environmental factors, and previous medical conditions (e.g. diabetes) are still

controversial, although they most likely strongly influence the development of the disease

(reviewed in Fang et al., 2015; van Rheenen et al., 2016). The remaining ~10 % of cases are

familial and are caused by specific mutations (Chen et al., 2013; reviewed in Renton et al.,

2014). The most common variations associated with ALS are within the genes of Chromosome

9 open reading frame 72 (C9orf72, ~30 %, DeJesus-Hernandez et al., 2011), SOD1 (~20 %,

first ALS gene to be identified in 1993 by Rosen et al.), TAR DNA-Binding Protein 43 (TDP-43,

5 %, Sreedharan et al., 2008), and fused in sarcoma (FUS, 5 %, (Kwiatkowski et al., 2009;

Vance et al., 2009; Naumann et al., 2019).

1.1.1 ALS-associated FUS

The FUS gene, also called translocated in liposarcoma, is located on chromosome 16 and

consists of 15 exons that encode a 526-amino-acid protein (Aman et al., 1996) and has first

been identified as a proto-oncogene causing liposarcoma by chromosomal translocation (Crozat

et al., 1993). It belongs to the FET protein family (the other members of that family being the

Ewing Sarcoma protein and TATA binding associated factor15, Svetoni et al., 2016), is

ubiquitously expressed in most tissues and mainly localizes to the nucleus (Andersson et al.,

2008; Sama et al., 2014). Its N-terminus is involved in transcriptional regulation and DNA

damage repair via a prion-like domain (rich in glutamine, glycine, serine, and tyrosine (QGSY)),

an additional glycine-rich region, and an RNA-recognition motif (Figure 1) (Sama et al., 2014).

The C-terminal region contains multiple arginine-glycine-glycine(RGG)-rich nucleic acid-binding

domains, a zinc-finger-binding domain, and a non-classical nuclear localization sequence

(Prasad et al., 1994) and attributes to e.g. RNA transport.



Figure 1: Functional domains of FUS.

The diversity of functional domains within FUS allows for its involvement in a multitude of cellular processes. Depicted are known domains of FUS and their annotated functions, as well as the most common mutations associated with familial ALS (adapted from Ticozzi et al., 2009; Sama et al., 2014; An et al., 2019). The mutation of interest in this study (P525L) is highlighted in red.

Due to this conserved, albeit complex structure, FUS interacts with DNA (Baechtold et al.,

1999) as well as RNA (Crozat et al., 1993; Zinszner et al., 1997) and its role in a variety of

cellular processes has long been proven. These include cell proliferation and DNA repair

(Bertrand et al., 1999), transcription regulation, RNA splicing (Yang et al., 1998) and transport

within the cell (Zinszner et al., 1997). FUS has been reported to bind RNA of more than 5500

genes in the central nervous system (CNS), primarily via a GUGGU-binding motif (Lagier-

Tourenne et al., 2012). By transporting mRNA to the distal axon or dendrites, e.g. those

encoding actin-related proteins, it may regulate local translation in response to external stimuli

(Fujii and Takumi, 2005; Fujii et al., 2005). Due to its diverse functions, it appears reasonable

that a series of pathological events are observed upon in its absence or malfunction.

1.1.2 Molecular pathology of ALS-FUS

Because FUS is involved in a variety of often interconnected processes within the cell, single

mutations in its genomic sequence can result in the occurrence of ALS (Figure 1), although it is

not yet clear whether the disease is caused by a loss-of-function or gain-of-function in FUS.

Mutations in FUS account for 35 % of cases manifesting before 40 years, typically presenting

classical ALS symptoms, and are responsible for the youngest case described (Conte et al.,

2012; Millecamps et al., 2012). These are autosomal-dominant mutations and primarily occur in

the C-terminal NLS, such as R521C or P525L, resulting in cytoplasmic mislocalization of FUS

due to reduced interaction with the nuclear import receptor Transportin-1 (Dormann et al., 2010)



and in impaired interaction of FUS with other RNA-binding proteins (RBPs) (Marrone et al.,

2019). Interestingly, there seems to be a direct relationship between the degree to which

nuclear import of FUS is diminished and the severity of the disease (defined by age of onset

and time till death) (Dormann et al., 2010; Kino et al., 2011; Niu et al., 2012).

FUS preferentially localizes to active chromatin (Immanuel et al., 1995), where it binds to

single-stranded DNA motifs in the promotor region of its target genes. Current evidence points

towards its function as a general transcriptional regulator, with only modest changes in mRNA

levels in its absence (Tan and Manley, 2010; Hoell et al., 2011; Lagier-Tourenne et al., 2012;

Rogelj et al., 2012; Tan et al., 2012; Li et al., 2013; Nakaya et al., 2013). FUS may also directly

bind to and recruit RNA polymerase II and III by preventing its phosphorylation at specific sites

(Immanuel et al., 1995; Bertolotti et al., 1996, 1998; Tan and Manley, 2010; Schwartz et al.,

2015). Additionally, FUS regulates transcription via interaction with specific transcription factors

(e.g. Runx2 or NF-κB, Du et al., 2011; Hallier et al., 1998; Kim et al., 2010; S. Sato et al., 2005;

Uranishi et al., 2001) and transcription initiation factor TFIID (Bertolotti et al., 1996; Hallier et al.,

1998; Uranishi et al., 2001; Li et al., 2010).

FUS modulates splicing of pre-mRNA in conjunction with other splicing regulators

machinery (e.g. SMN, U11, U11, U12, Sm proteins, SFPQ, and others) (Alliegro and Alliegro,

1996; Hackl and Lührmann, 1996; Zhou et al., 2002; Meissner et al., 2003; Yamazaki et al.,

2012; Gerbino et al., 2013; Tsuiji et al., 2013; Sun et al., 2015; Reber et al., 2016; Ishigaki and

Sobue, 2018) by binding to long introns within the prespliced RNA (Hoell et al., 2011; Ishigaki et

al., 2012; Lagier-Tourenne et al., 2012; Rogelj et al., 2012). As such, FUS is involved in the

splicing of actin pre-mRNA, as has been shown for example in mouse brain extracts (Fujii and

Takumi, 2005), but also of the microtubule-associated protein (MAP) tau, a neuron specific

cytoskeleton related protein (Ishigaki et al., 2012; Lagier-Tourenne et al., 2012; Orozco et al.,

2012; Rogelj et al., 2012). Altered splicing by mutant FUS variants hence might affect the

cytoskeletal architecture of neurons. FUS also regulates its own splicing (Lagier-Tourenne et al.,

2012; Nakaya et al., 2013), specifically by FUS-mediated skipping of exon 7 in FUS pre-mRNA,

which results in its nonsense-mediated decay (Ishigaki and Sobue, 2018). This process is

disrupted for mutant FUS variants due to their mislocalization to the cytoplasm. Impaired FUS

splicing may in turn exacerbate the pathogenic cytoplasmic accumulation of FUS (Zhou et al.,

2013). Further, FUS is involved in alternative splicing of more than 3200 exons, of which many

are part of mRNAs coding for proteins important in neuronal function or neurodegeneration

(Ishigaki et al., 2012).



There is evidence that the physical binding capability between FUS and its RNAs is not

altered between wildtype and mutant FUS variants, as 80 % of transcripts binding to mutant

(R521G and R521H) FUS can also bind to wildtype FUS (Hoell et al., 2011). This supports the

hypothesis of a gain-of- function phenotype for mislocalized cytoplasmic mutant FUS variants

with respect to RNA binding and processing. After splicing, FUS subsequently shuttles mRNA

between the nucleus and the cytoplasm (Fujii et al., 2005). FUS has been attributed to interact

with several actin- and microtubule-binding motor proteins, including Myosin-Va (Yoshimura et

al., 2006) as well as Myosin-VI (Takarada et al., 2009), and has been isolated as part of large

granules that associates with KIF5B (Kanai et al., 2004). This hints towards an involvement of

FUS in cellular transport of mRNAs to their dedicated locations for local translation. Indeed,

upon synaptic activation via the metabotropic glutamate receptor mGluR5, FUS translocates

into dendritic spines (by to date unknown mechanisms) and may facilitate the local translation of

actin-associated proteins (Fujii and Takumi, 2005). This is in line with the fact that loss of FUS

consequently results in abnormal spine morphology and attenuated spine density (Fujii et al.,

2005), since spine structure and stability is largely dependent on the actin cytoskeleton (Penzes

and Rafalovich, 2012; Basu and Lamprecht, 2018). Local translation is however not solely

regulated by the transport of mRNA, but also by sequestering mRNA together with its RBPs.

As a protein with primarily nuclear localization under physiological, non-stressed

conditions, FUS binds single- and double-stranded DNA (Baechtold et al., 1999; Liu et al.,

2013), thereby promoting DNA damage repair via homologous recombination (HR) and

nonhomologous end joining (NHEJ) (Mastrocola et al., 2013; Wang et al., 2013). For HR, the

pairing of homologous DNA is an essential step which has been proposed to be a crucial

function devoted especially to the C-terminal region of FUS (Akhmedov et al., 1995; Baechtold

et al., 1999; Liu et al., 2013). Phosphorylated FUS (Pezzano et al., 1996) is one of the first

proteins recruited to sites of DNA damage (Mastrocola et al., 2013; Wang et al., 2013), where it

interacts with histone deacetylase 1 (HDAC1) and poly-ADP ribose (Deng et al., 2014). A

subset of ALS-associated mutations impair FUS in its function in DNA damage response

(Rulten et al., 2014; Naumann et al., 2018). In the absence or in case of malfunction of FUS,

DNA double-strand repair by HR and NHEJ was decreased between 30-50 % (Mastrocola et al.,

2013; Wang et al., 2013). In primary mouse cortical neurons, the decrease in DNA repair

efficiency by NHEJ upon FUS depletion was even higher, namely 65 % (Wang et al., 2013).

Together, these results suggest a role of FUS in DNA damage repair in proliferating as well as

postmitotic cells. It has further been shown that FUS interacts with HDAC1 at sites of DNA

double-strand breaks (Miller et al., 2010; Dobbin et al., 2013; Thurn et al., 2013), while it is



recruited to single-strand breaks by poly adenosine diphosphate ribose polymerase (PARP),

which binds to these sites and subsequently polymerizes poly adenosine diphosphate ribose

(PAR) chains (Schreiber et al., 2006; De Vos et al., 2012). In a transgenic mouse model

expressing FUS-R521C next to its wildtype form, mutant FUS forms a complex with wildtype

FUS and alters its interaction with HDAC1 (Qiu et al., 2014), calling for a dominant negative

effect of this FUS mutation. This is in line with the finding that several mutant FUS variants

(R244C, R514S, H517Q, and R521C) showed deficiencies in HR--mediated repair relative to

wildtype FUS in U2OS cells, independent of their nuclear or cytoplasmic localization (Wang et

al., 2013). As DNA damage accumulates with increasing age (e.g. due to a lifetime exposure to

DNA-damaging factors and a decay of quality control mechanisms over decades, Gorbunova et

al., 2007), it seems logical that the pathological consequences, i.e. the phenotype of FUS -ALS,

becomes apparent only later in the life of patients. It also indicates why mainly neurons are

affected by the disease, since they lack the ability to replicate and self-renew and are therefore

more susceptible to accumulate DNA damage. This is in line with the finding of increased levels

of γH2AX in postmortem brain sections of patients harboring the FUS R521C or P525L mutation

(Wang et al., 2013). γH2AX is a marker for DNA damage, but also correlates with apoptosis

(Rogakou et al., 2000; Pasinelli and Brown, 2006).

The low complexity, prion-like domain of FUS, with its high content of glycine, accounts for

its tendency to aggregate (Han et al., 2012). Under physiological conditions, RBPs with such a

domain can accumulate together with mRNA, ribosome translation initiation factors, and other

proteins into membraneless compartments called stress granules (Dewey et al., 2012; Mateju et

al., 2017). Stress granules are stalled translational complexes evolving upon metabolic or

environmental stress involved in the triage of mRNAs, determining whether an mRNA is

translated, degraded, or suppressed in order to promote the expression of proteins essential to

reestablish homeostasis (Sama et al., 2014). Wildtype FUS can rapidly and reversibly shuttle to

the cytoplasm upon induction of cellular stress, where it associates with stress granules. Under

physiological conditions, it returns to the nucleus once stress is resolved (Dormann et al., 2010).

ALS-associated, consistently mislocalized FUS variants, however, have long been observed to

assemble within pathological stress granules upon protein overexpression, heat shock, and ER

or oxidative stress (Andersson et al., 2008; Bosco et al., 2010; Dormann et al., 2010; Gal et al.,

2011; Kino et al., 2011; Bentmann et al., 2012; Daigle et al., 2013). However, if mutated variants

of RBPs are incorporated into these structures, they undergo a pathological liquid-to-solid phase

transition, resulting in solidified SGs (Bowden and Dormann, 2016). Solidified SGs have lost

their dynamicity and show altered mechanical properties (Nötzel et al., 2018), and hence can no



longer fulfill their physiological roles. This results in impaired stress response and mRNA

transport, changes in local translation, and the formation of pathological aggregates, all of which

contributes to neuron dysfunction and the progression of neurodegeneration (Bowden and

Dormann, 2016). The formation of SGs and how severely it affects the cell therefore also largely

depends on the local concentration of a respective protein. For instance, cytoplasmic

accumulated mutant FUS colocalizes with these pathological aggregates (Dewey et al., 2012;

Wolozin, 2014). Hence, while FUS itself is not essential for the formation of SGs, the

cytoplasmic mislocalization of its mutant variants largely contributes to the transformation of

SGs.

Another class of cellular organelles that seems to be substantially influenced by mutant

FUS variants is mitochondria. Disruptions in various mitochondrial characteristics, such as

structure, dynamics, bioenergetics, calcium homeostasis, endoplasmatic reticulum (ER)-

mitochondrial contact, mitochondrial quality control, and cellular transport have been reported in

ALS patients and various model systems (reviewed in Smith et al., 2019). One of the first

observed changes in ALS patient motor neurons are structurally altered (i.e. swollen and

vacuolated) and aggregated mitochondria (Atsumi, 1981; Sasaki and Iwata, 2007).

Overexpression of FUS R521G/H or P525L variants results in mitochondrial shortening

(Tradewell et al., 2012; Deng et al., 2015; Sharma et al., 2016). The P525L mutation additionally

results in deformation and loss of mitochondrial cristae in mouse models (Deng et al., 2015;

Sharma et al., 2016). FUS seems to directly influence mitochondrial function through interaction

with the mitochondrial chaperone heat shock protein of 60 kDa (HSP60) (Deng et al., 2015) and

overexpression of FUS leads to reduced mitochondrial ATP production (Stoica et al., 2016) as

well as augmented levels of reactive oxygen species (ROS), causing oxidative stress (Deng et

al., 2015).

Non-functional and damaged mitochondria are subject to clearance by mitophagy under

physiological conditions and need to be replaced by functional mitochondria (Sterky et al., 2011;

Hamacher-Brady and Brady, 2016). This process requires a robust transport of mitochondria

throughout the cell. However, in motor neurons carrying the P525L mutation, generated from

ALS patient-derived induced pluripotent stem cells (iPSCs), virtual arrest of mitochondrial

movement in distal, but not proximal axons was observed, accompanied by a loss of membrane

potential in these mitochondria along with reduced mitochondrial length (Naumann et al., 2018).

In general, a defect in transport of mitochondrial and other organelles has reportedly been

observed, often as one of the earliest pathophysiological events in motor neurons affected by



ALS, which indicates that deficits in axonal transport might be a primary cause of motor neurons

loss (De Vos et al., 2008; De Vos and Hafezparast, 2017a).

1.2 Axonal transport

Intracellular transport of RNA, proteins, and organelles is essential to cell survival and correct

function, as well as neurotrophic and injury signaling. It becomes particularly important in

(motor) neurons, which have to facilitate long-range transport along dendrites and axons, latter

with lengths up to one meter (Grafstein and Forman, 1980; Fletcher and Theriot, 2004). In

addition to the large distance that needs to be covered by cellular transport mechanisms,

cellular homeostasis requires transport in two directions, namely anterograde (i.e. away from

the cell soma) and retrograde (i.e. towards the cell soma). Two main classes of axonal transport

can be distinguished based on the average speed of transport movement. Fast axonal transport

is characterized by a speed of ~50 - 400 mm/day or 0.6 - 5 µm/s. Slow axonal transport is

further subdivided into two parts, depending on the proteins transported and the speed, namely

slow component a and b, which cover a distance of 0.2 - 3 mm/day or 0.0002 - 0.03 µm/s and

2 - 10 mm/day or 0.02 - 0.1 µm/s, respectively (De Vos and Hafezparast, 2017a). Each of these

transport types delivers a distinct set of cargoes, and the differences in overall speed results

from prolonged pauses between movement phases in slow axonal transport (Black, 2016). Both

fast and slow transport is mediated by the same molecular motors moving along tracks of the

cytoskeletal network, defining motors and the cytoskeleton as the two major components of

axonal transport.

1.2.1 Microtubules as part of the neuronal cytoskeleton

The neuronal cytoskeleton consists of three major components, namely actin filaments,

neurofilaments, and microtubules. Actin filaments are composed of granular (G-) actin

monomers, which assemble to form a ~6 nm wide double-helix of filamentous (F-) actin (Boron

and Boulpaep, 2005) and are most prominent in the axonal growth cone and dendritic spines

(Schevzov et al., 2012). Neurofilaments are type IV intermediate filaments with an average

diameter of ~10 nm and are present in the perikarya and dendrites, but particularly in axons,

where they are vital for the radial growth of axons during development and the further

maintenance of axon caliber, as well as the transmission of electrical impulses along the axon

(Friede and Samorajski, 1970; Ohara et al., 1993; Eyer and Peterson, 1994; Zhu et al., 1997;

Yum et al., 2009; Yuan et al., 2012).

The major portion of axonal transport occurs along microtubules, which are rigid hollow

cylinders about 25 nm in diameter (Figure 2). They are dynamic filaments that frequently



assemble and disassemble within the cell. Microtubules polymerize from free tubulin, which

itself is a heterodimer of two closely related 55 kDa polypeptides, α- and β-tubulin. Microtubules

are formed by 13 linear chains of these heterodimers, so called protofilaments. Both tubulin

polypeptides bind GTP to regulate polymerization, but only GTP bound to β-tubulin is

hydrolyzed shortly after polymerization. GTP hydrolysis weakens the binding affinity of β-tubulin

to adjacent molecules, which favors depolymerization and enables dynamic behavior of

microtubules (Cooper, 2000a). This behavior is referred to as dynamic instability and describes

the alternating cycles of growth and shrinkage of microtubules at their individual protofilaments,

a process determined by the rate of GTP hydrolysis relative to the rate of tubulin dimer addition

(Mitchison and Kirschner, 1984). Since the sequential addition of tubulin heterodimers results in

so called head-to-tail arrays (Cooper, 2000b) with alternating α- and β-tubulin units in each

protofilament, microtubules are polar structures with a fast-growing plus end (protofilament

ending with a β-tubulin subunit) directed towards the cell cortex, and a slow-growing, more

stable minus end (protofilament ending with a α-tubulin subunit) directed towards the soma

(Maday et al., 2014). In axons, microtubules form a unipolar array with the plus end facing

towards the distal axon, while the microtubule organization in dendrites often consists of arrays

with mixed polarity (Baas et al., 1988; Kwan et al., 2008; Kleele et al., 2014). Hence, the correct

polarity and orientation of microtubules majorly contribute to the specific distribution of cargo

throughout the cell by a variety of motor proteins.



Figure 2: The motor protein kinesin-1 transports cargo along microtubules.

Microtubules are hollow tubes with a diameter of 25 nm, which are composed of 13 individual protofilaments formed by the continuous addition of α- and β-tubulin heterodimers. GTP bound to β-tubulin becomes rapidly hydrolyzed after polymerization, which weakens the binding affinity of tubulin to adjacent molecules and allows for depolymerization of the microtubule, a continuous process referred to as dynamic instability. Microtubule growth occurs if GTP-bound tubulin is added more rapidly to the growing microtubule than GTP is hydrolyzed, with the rate of polymerization being typically much faster at the plus end of the microtubule, where β-tubulin is exposed, than at the minus end, where α-tubulin is exposed. Microtubules are stabilized by microtubule associated proteins such as tau, which on the other hand also act as roadblocks for microtubule-associated motors walking along a protofilament, hindering their motility. This figure shows the microtubule-associated motor kinesin-1, a heterotetramer consisting of two kinesin heavy chains and two kinesin light chains (KLC). One kinesin heavy chain is comprised of a catalytic motor domain, which hydrolysis ATP to facilitate movement in 8 nm steps across the microtubule, a neck linker region, an α-helical stalk with two hinge regions required for dimerization, and a tail domain interacting with KLCs to mediate cargo binding (Cooper, 2000a).



1.2.3 Microtubule motor proteins

There are two major families of motor proteins that associate with microtubules, namely kinesins

and dyneins. The human kinesin superfamily consists of 45 members, of which 38 are

expressed in the brain (Miki et al., 2001), and can be subdivided into 15 subfamilies. Most

kinesins move towards the plus end of microtubules in a straight path along a single

protofilament (Maday et al., 2014), thereby mainly driving anterograde transport in neuronal

axons. Members of the kinesin-1 (also referred to as KIF5), kinesin-2 (i.e. KIF3A and B, KIF17),

and kinesin-3 (i.e. KIF1A, KIF1Bα, and KIF1Bβ) family collectively contribute to this anterograde

(Maday et al., 2014), with fast axonal transport mainly involving kinesin-1 and members of the

kinesin-3 family, while slow anterograde axonal transport appears to be mainly mediated by

kinesin-1 (Xia et al., 2003). Kinesin-1 is to date the best studied motor protein and was first

discovered in axons of the giant squid in 1985 (Vale et al., 1985). It is a heterotetramer

consisting of two kinesin heavy chains (KHCs), encoded by the three mammalian genes KIF5A,

B, and C, and in most cases two kinesin light chains (KLCs), which are substantial for cargo

binding and in part contribute to the autoinhibition of kinesin in the absence of cargo (Figure 2)

(Sun et al., 2011a). KHCs are comprised of the catalytic motor domain, which facilitates walking

along the microtubule by hydrolyzing ATP and conveys processivity and direction of movement

together with a neck linker region. The motor domain is linked to an α-helical stalk interrupted by

two hinge regions, which is required for dimerization. Further, a tail domain interacts with cargo

either on its own or through KLCs and facilitates autoinhibition of the motor domain (Williams et

al., 2014, reviewed in Nobutaka Hirokawa et al., 2010). Cargo binding often requires a variety of

adaptor proteins, such as c-Jun N-terminal kinase-interacting protein (JIP), trafficking kinase

(TRAK), or mitochondrial Rho GTPase (Miro) 1 and 2, which directly or indirectly (i.e. via KLCs)

link kinesin-1 to specific cargoes (Fu and Holzbaur, 2014). Kinesin-1 walks in a hand-over-hand

mechanism with a step size of ~8 nm (about the size of a tubulin dimer) for approximately 100

steps before it detaches, walks at a speed of ~0.5 - 1 µm/s and a stall force of 5 - 6 pN

(Svoboda and Block, 1994; Hackney, 1995; Hua et al., 1997; Coy et al., 1999; Yildiz et al.,

2004; Asbury, 2005) Cargoes transported by kinesin-1 include organelles such as mitochondria,

as well as vesicular and non-vesicular cargoes (i.e. lysosomes, signaling endosomes containing

e.g. brain-derived neurotrophic factor (BDNF), amyloid precursor protein vesicles, AMPA

vesicles, and mRNA/protein complexes (Hirokawa et al., 2010).

In contrast, members of the kinesin-2 family can assemble into both homodimeric and

heterodimeric motors (Scholey, 2013) and transport for example fodrin-positive plasma

membrane precursors (Takeda et al., 2000), N-cadherin and β-catenin (Teng et al., 2005),



choline acetyltransferase (Ray et al., 1999), and Rab7-positive late endo-lysosomes (Hendricks

et al., 2010; Castle et al., 2014). Members of the kinesin-3 family dimerize upon cargo binding to

form highly processive motors (Soppina et al., 2014) and drive the motility of dense core

vesicles and synaptic vesicle precursors (Hall and Hedgecock, 1991; Okada et al., 1995; Lo et

al., 2011).

While anterograde transport is driven by a pool of different kinesins, retrograde transport

is almost exclusively carried out by cytoplasmic dynein (Roberts et al., 2013). Cytoplasmic

dynein belongs to the family of AAA+ ATPases and can be subdivided into cytoplasmic dynein 1

and 2. Cytoplasmic dynein 1 (hereafter referred to as dynein) is the main retrograde molecular

motor in neurons. It is a large, ~1.5 kDa protein complex comprising two homodimeric dynein

heavy chains (DHCs) dimerizing by their N-terminal tail domain, and multiple dynein

intermediate chains, dynein light intermediate chains, and light chains (King, 2012). The latter

three are encoded by several genes giving rise to various isoforms, and therefore contribute to

cargo-specific recruitment (Kuta et al., 2010). Similar to kinesin-1, multiple adaptor proteins,

including the dynactin complex, Bicaudal D2, lissencephaly 1, or nuclear distribution protein,

regulate the correct function, specific cargo binding and localization of dynein (Kardon and Vale,

2009). Dynein is a fast motor with velocities ranging from 0.5 - 1 µm/s, but unlike kinesin, it

frequently takes back- and side-steps between microtubule protofilaments (Mallik et al., 2005;

Ross et al., 2006). The coordinated activity of multiple motor or the binding of activators such as

Bicaudal D2, however, converts dynein into a unidirectional, highly processive motor (Mallik et

al., 2005; McKenney et al., 2014; Schlager et al., 2014). With ~1 pN, the stall force of dynein is

much weaker than those of kinesins (e.g. 5-7 pN for processive kinesin-1 or 1.5-7 pN for weakly

processive kinesin-5 in vitro, Hesse et al., 2013; Schroeder et al., 2010).

Most cargoes are not transported by a single type of motor, but by an ensemble of various

oppositely directed kinesins and dyneins (Hendricks et al., 2010; Encalada et al., 2011), even if

they are processively transported in a single direction over long distances (Maday et al., 2012).

A single organelle may be transported by 1-2 kinesins and 6-12 dyneins (Hendricks et al., 2010;

Schroeder et al., 2012; Rai et al., 2013). Several models have been suggested as to how this

assembly of motors is organized (Gross, 2004; Welte, 2004; Gross et al., 2007). The simplest

model postulates an unregulated tug-of-war between opposing kinesins and dyneins, which

could be successfully modelled for late endo-lysosomes (Müller et al., 2008; Hendricks et al.,

2010). In this scenario, movement-of-directionality of cargoes might be determined by the

biophysical characteristics of motors themselves. Kinesin-2 for instance shows load

force-dependent attachment, indicating it may be less likely to win in a tug-of-war situation



(Schroeder et al., 2012). In contrast, other cargoes such as autophagosomes exhibit strong

unidirectional motility (Maday et al., 2012), suggesting a possible down-regulation of kinesin and

pointing towards two alternative models: i) the coordinated regulation of all motor types on a

single cargo, so that only one type of motor is active at a given time, or ii) a tight regulation of

only one type of motor, i.e. kinesin as it is the stronger motor, while the other, i.e. the weaker

dynein motor, might simply be overpowered if both motors are active simultaneously and only

becomes active when the stronger motor is downregulated. This is supported by evidence for a

regulatory role of scaffolding proteins to regulate opposing motors on the same cargo (Fu and

Holzbaur, 2014; Fu et al., 2014). In addition, the well-studied autoinhibitory mechanisms of

kinesin-1 provides further evidence for this model: in the absence of cargo, the kinesin-1 tail

domain binds to the motor domain and thereby blocks motor function (Kaan et al., 2011), which

is only relieved by binding of specific scaffolding proteins such as JIP1 and JIP3 (Blasius et al.,

2007; Sun et al., 2011a). Apart from kinesin-1, regulatory mechanisms for other kinesins or for

dynein have not been elucidated in such detail to date.

The exact regulatory mechanism for each cargo most likely results from combinations of

these models. Nevertheless, a few general commonalities can be found among the diverse

patterns of motility: i) during transport, motors remain stably associated with their cargo, even in

their inactive state; ii) cargo can be effectively transported over large distances (>1 µm) by a

small ensemble of (usually opposing) motors; iii) regulatory mechanisms to control motor activity

include specific recruitment by Rab-GTPases, scaffolding proteins, and upstream regulation by

kinases and phosphatases (Maday et al., 2014).

As motor proteins are often biased, if not limited, by their preference of walking towards

one or the other end of a microtubule, they are highly influenced by the assortment of MAPs and

other posttranslational modifications they encounter along their way. While MAPs often act as

physical barriers for motors walking on cytoskeletal tracks, they can also alter the stability of

these tracks and thereby influence motor motility.

1.2.2 Microtubule-associated protein tau

Because of their dynamic instability, microtubules frequently disassemble and assemble within

the cell. Disassembly is mediated either by microtubule severing proteins or by increasing the

rate of tubulin depolymerization at the microtubule ends. Continuous assembly, i.e. the growth

of microtubules, requires stabilization of joined tubulin heterodimers, which can be achieved

through posttranslational modifications of tubulin, such as detyrosination or acetylation, or

through tightly regulated interactions with MAPs. The best characterized MAPs are MAP-4 in



non-neuronal cells, and MAP-1, MAP-2 (which is absent in axons), and tau (which is absent in

dendrites, but abundant in axons) in neuronal cells (Weingarten et al., 1975; Witman et al.,

1976; Cooper, 2000a). The ability of MAPs to interact with microtubules highly depends on the

posttranslational modifications of MAPs, specifically phosphorylation in case of tau (Lindwall

and Cole, 1984; Mandelkow et al., 1995; Drewes et al., 1997; Stoothoff and Johnson, 2005;

Ramkumar et al., 2018). Hyperphosphorylation of tau is associated with a variety of diseases

(Ksiezak-Reding et al., 1992; Köpke et al., 1993), but also with fetal development (Matsuo et al.,

1994; Yu et al., 2009) or a state of hibernation (Arendt et al., 2003). Tau is to date the best

studied MAP, in part because of its implication in a group of neurodegenerative diseases called

tauopathies (e.g. AD) (Brion et al., 1986).

The tau gene is located on chromosome 17q21 and is comprised of 16 exons, of which

exon 2,3, and 10 are alternatively spliced to produce six different isoforms in the human brain

(Goedert et al., 1989a, 1989b). Exons 2 and 3 encode for two amino-terminal inserts (N), while

exon 10 encodes for one of a total four microtubule-binding repeats (R), followed by a short C-

terminal tail sequence (Schweers et al., 1994). An isoform hence contains 0N, 1N or 2N as well

as 3R or 4R and ranges from 352 to 441 amino acids (45-75 kDa) in length (Figure 3) (Bukar

Maina et al., 2016; Guo et al., 2017a; Pîrşcoveanu et al., 2017). The isoform 0N3R is the

predominant isoform expressed during fetal development, while all isoforms are expressed in

the adult brain (Goedert et al., 1989b; Hefti et al., 2018).The C-terminal microtubule-binding

repeats form the microtubule-binding domain of tau, with their in- or exclusion influencing the

tau microtubule binding affinity, while the N-terminal domain projects away from the microtubule

surface, possibly determining the space between microtubules and interacting with other

cytoskeletal components (Hirokawa et al., 1988; Chen et al., 1992; Lu and Kosik, 2001). Under

physiological conditions, tubulin is present in 10- to 20-fold molar excess of tau in mature adult

neurons, with a physiological concentration of tau ~2-4 µM (Butner and Kirschner, 1991;

Khatoon et al., 1992; Dixit et al., 2008; Avila, 2010) and a ratio of 3R to 4R isoforms of about 1:1

(Goedert et al., 1989a; Andreadis, 2005; Ishigaki et al., 2017; Pîrşcoveanu et al., 2017).

Interestingly, recent publications demonstrated an increase in 4R:3R ratio upon knockdown of

FUS and demonstrate that FUS, in combination with splicing factor proline- and glutamine-rich

(SFPQ), directly regulates the splicing of tau mRNA (Ishigaki et al., 2017), pointing towards an

involvement of tau isoform ratio changes in FUS-ALS pathology.



Figure 3: Functional domains of tau.

Six different isoforms of tau are expressed in the adult human brain. Since there is only one gene coding for tau, isoforms are created by alternative splicing of exons 2 and 3, coding for one of two N-terminal repeats (N1 or N2), and exon 10, coding for the second of four microtubule binding repeats (R2). The projection domain, facing away from the microtubule surface, is linked to the microtubule binding domain through a proline-rich region. Depicted are known domains of tau and their annotated functions (adapted from Ambadipudi et al., 2017; Bukar Maina et al., 2016; T. Guo et al., 2017; Polverino de Laureto et al., 2019; Tepper et al., 2014).

The constant association and disassociation of tau with microtubules, along with the

different microtubule binding affinities of each tau isoform, provides microtubule stabilization as

well as flexibility as needed for correct cytoskeletal organization, especially in the distal part of

the axon (Black et al., 1996; Kempf et al., 1996). Tau localizes to labile, unstable domains of

microtubules (Black et al., 1996), where it increases its stability, thereby promoting assembly

while preventing depolymerization (Brandt and Lee, 1993; Panda et al., 1995). On the other

hand, MAPs bound to microtubules may act as roadblocks for microtubule-associated motor

proteins walking along individual protofilaments. Especially kinesin-1 has been shown to detach

at patches of microtubule-bound tau, while dynein tends to reverse direction upon encountering

lattice bound tau (Dixit et al., 2008). In addition, kinesin-1 has been proposed to compete with

tau for the same binding site on microtubules (Stamer et al., 2002; Dubey et al., 2008).

The sensitive interplay between motors, cargoes, microtubules and their posttranslational

modifications as described above can be disturbed, leading to perturbed axonal transport

perturbed as observed in for example ALS.



1.2.4 Axonal transport in ALS

First evidence for defects in axonal transport as part of ALS pathology came from post mortem

studies that reported abnormal accumulations of cytoskeletal filaments and organelles, such as

mitochondria and lysosomes (Hirano et al., 1984b, 1984a; Rouleau et al., 1996). Since then,

numerous studies have been carried out on endosome and mitochondrial trafficking or the

distribution of mRNA-containing granules (reviewed in (De Vos and Hafezparast, 2017a),

highlighting the importance of axonal transport defects in ALS pathology as a pathological event

preceding symptoms of ALS in a mouse model (Bilsland et al., 2010). However, most studies

focused on axonal transport defects associated with mutant SOD1 variants. Only little is known

about the extent and underlying pathological mechanisms of axonal transport defects caused by

abnormal FUS variants. The FUS-P525L variant causes severe impairment in axonal transport

of mitochondria in a Drosophila melanogaster model of ALS and in spinal motor neurons

differentiated from ALS patient-derived iPSCs (Baldwin et al., 2016; Naumann et al., 2018).

FUS has been proven to directly bind and regulate expression of mRNAs for several

motor proteins such as KIF5C and KIF1B (Guo et al., 2020), which are part of the ensemble that

regulates mitochondrial movement and distribution in neurons (Schwarz, 2013; Campbell et al.,

2014). In addition, stress granules undergoing a pathological liquid-to-solid phase transition

caused by FUS-NLS mutant variants, as described above (see 1.1.2), sequester RNA and

proteins in the cytoplasm, including those of the axonal transport machinery (Aulas and Vande

Velde, 2015). A more indirect influence of FUS on axonal transport has been proposed due to

the fact that expression levels of histone deacetylase 6 (HDAC6) are decreased when FUS is

silenced in mammalian cells, leading to cytoskeletal changes due to aberrant acetylation and

hence ultra-stabilization of microtubules (Hubbert et al., 2002; Kim et al., 2010). Conversely,

Guo et. al. could show that axonal transport is restored in FUS-ALS patient-specific

iPSC-derived motor neurons upon HDAC6 inhibition (Guo et al., 2017b), suggesting that the

role of tubulin acetylation in causing axonal transport defects is more complex and needs further

investigation. Changes in the underlying cytoskeletal tracks likely influence the motility of motors

walking on them. Apart from impaired transport itself, improper functioning of mitochondria has

also been suggested to contribute to ALS pathophysiology. Several FUS variants with a

mutation in their NLS disrupt mitochondrial ATP production and calcium homeostasis. The latter

is likely caused by dysfunctional communication between the ER and mitochondria due to a

reduction of their contact sites (Stoica et al., 2016). A disruption in mitochondrial ATP production

might indicate that impaired transport is due to a local depletion of ATP, the essential energy

source for motor motility.



Our limited understanding of the pathomechanisms involved in axonal transport defects in

FUS-ALS highlight the importance for further studies addressing this matter. A number of model

systems have been established that mimic FUS-ALS pathology and/or axonal transport in

general.

1.3 Modelling ALS and axonal transport in vitro

Age-related NDs are human-specific, as non-human primates or rodents do not readily develop

comparable neuropathological or clinical phenotypes (Mullane and Williams, 2019). Despite this

fact, especially mouse and rat models have long been the gold standard to study NDs in general

(Petrov et al., 2017; Van Damme et al., 2017), and although the results obtained from these

studies have been fundamental in uncovering disease pathogenesis, potential drugs identified in

animal trials have failed in clinical tests (Matus et al., 2014).

1.3.1 FUS-ALS specific phenotypes in iPSC-derived motor neurons

A more physiological model system for NDs developed with the availability of iPSCs. iPSCs are

generated by transfecting e.g. the so called Yamanaka transcription factors (Oct3/4, Sox2, Klf4,

and c-Myc) into human adult somatic cells, which induce the reprogramming and conversion of

these somatic cells to pluripotent stem cells (Takahashi and Yamanaka, 2006; Takahashi et al.,

2007). This technique allows for the generation of patient-specific iPSCs from e.g. human

fibroblasts, which are easily obtained by a skin biopsy. After successful regeneration, iPSCs can

be differentiated into virtually every cell type (Yu et al., 2007; Aasen et al., 2008), including

neuronal subtypes (Emdad et al., 2012; Japtok et al., 2015; Maury et al., 2015; Douvaras et al.,

2016), and have as such been used to model ALS (Chen et al., 2014; Kiskinis et al., 2014;

Wainger et al., 2014; Japtok et al., 2015). One of the most important advantages of this model

system is that neurons, or every other cell type, derived from patient-specific iPSCs exhibit

endogenous pathomechanisms. Such might be masked in conventional transgenic models due

to unphysiological overexpression of proteins, species-specific differences in protein expression

profiles, or the lack of availability of cell types of interest (e.g. human postmitotic neurons). The

emergence of improved gene-engineering techniques, such as CRISPR/Cas9, further increased

the capability of iPSC models to study intrinsic and endogenous cellular mechanisms, as it

allows for the reversion of single mutations to their wildtype state or the introduction of a specific

mutation into the wildtype genome (Deveau et al., 2010; Deltcheva et al., 2011; Jinek et al.,

2012; Hsu et al., 2013; Ran et al., 2013). Pairs of so called isogenic cell lines can thereby be

created from a single donor line, which contain either the wildtype or the mutated form of a

specific gene in an otherwise identical genetic background. Hence, differences observed



between those two lines can quite precisely be attributed to the individual and direct influence of

a genetic mutation.

Although motor neurons derived directly from iPSCs have been used in the past for drug

screening purposes (Egawa et al., 2012), the technique required further improvement as the

cultivation of iPSCs requires expensive growth factors, frequent feeding and splitting at narrow

ratios and needs significant time for differentiation, while it often results in inefficient

differentiation. Therefore, protocols were developed that prime iPSCs towards certain cell

lineages, e.g. neuronal cell types, using only a limited set of small molecules, i.e. growth factors

or inhibitors influencing WNT and sonic hedgehog signaling (Reinhardt et al., 2013). Using such

protocols, neuronal precursor cells (NPCs) can be derived from iPSCs, which are much easier

to propagate and can be efficiently and repeatedly differentiated into e.g. spinal motor neurons

using a different set of small molecules (Reinhardt et al., 2013; Naumann et al., 2018). Spinal

motor neurons differentiated from iPSC-derived NPCs of two isogenic cell lines can hence be

utilized to specifically and reproducibly investigate the direct influence of a single ALS-

associated mutation on cellular (patho-) mechanisms in an endogenous environment.

1.3.2 Reconstitution of axonal transport in vitro

In cell culture models, it is difficult to study the contribution of a single protein to pathological

changes in its endogenous environment. One particular protein is often involved in various

cellular pathways and in its absence or malfunctioning, it can in some cases be substituted by

other proteins. The involvement of this protein in a particular cellular mechanism might hence be

masked in cell culture models. To address this limitation, minimal systems have emerged to

study the direct interaction of a few much defined components. One of these systems is the so

called stepping assay, where single fluorescently labelled motors move across

surface-immobilized microtubules. First experimental setups involved binding of motors to

micron-sized beads, which can be tracked with nanometer precision (Howard, 2001), and

subsequent video imaging (Sheetz and Spudich, 1983; Yanagida et al., 1984; Spudich et al.,

1985) or optical trap experiments to measure forces generated by motors (Svoboda et al., 1993;

Mehta et al., 1999; Rief et al., 2000; Schnitzer et al., 2000). With advances in microscopy

techniques, e.g. TIRF microscopy, the use of smaller cyanine-based fluorophores, such as Cy3

(Funatsu et al., 1995; Vale et al., 1996), and GFP (Pierce et al., 1997) coupled to individual

kinesin molecules became possible. To date, these single molecule experiments have not only

been used to study the stepping characteristics of motors, but also (de-) polymerizing activities

and diffusion of motors and various MAPs (Yildiz et al., 2003; Helenius et al., 2006; Varga et al.,



2006, 2009; Bieling et al., 2007; Brouhard et al., 2008; Fink et al., 2009; Bugiel and Schäffer,

2018; Mitra et al., 2018).

Another method to reconstitute cellular transport in vitro has been developed with gliding

motility assays, where fluorescently labelled microtubules or actin filaments are propelled by

underlying surface-immobilized molecular motors, using ATP as an energy source. Motors are

typically immobilized on surfaces either via antibodies or by adsorption (Figure 4), while the

number of motors can vary and even be reduced to the point where a microtubule is propelled

by a single motor (Howard et al., 1989). The setup can be easily observed and analyzed using

conventional widefield fluorescence microscopy (Nitzsche et al., 2010). In the past, actin

filaments as narrow as 6 nm could observed by dark-field (Nagashima and Asakura, 1980) or

epifluorescence microscopy (Yanagida et al., 1984).

Figure 4: Principle of a microtubule gliding motility assay.

Fluorescently labeled microtubules are propelled by truncated kinesin-1 motors that are immobilized on a glass surface. As a first step in assays preparation, the assay’s glass surface is blocked with casein to prevent unspecific binding (Nitzsche et al., 2010).

By tracking the distance a microtubule filament is propelled by multiple underlying motors within

a defined time, the integrated motility of the underlying motors can be determined (Ruhnow et

al., 2011). The best understood and hence most widely used motor in these assays is kinesin-1,

but other motors can and have also been employed (Braun et al., 2011, 2017; Herold et al.,

2012; Monzon et al., 2019). Microtubules in vivo are characterized by high dynamic instability,

but often need to be highly stabilized for the use in microtubule gliding assays. This can be

achieved by growing them in the presence of the slowly hydrolysable GTP analogue guanylyl-

(α, β)-methylene-diphosphonate (GMP-CPP) or by adding microtubule stabilizing drugs such as



taxol after polymerization (Schiff et al., 1979; Korten et al., 2010), a drug which has also been

used for in vivo applications.

Taxol, or paclitaxel, was first isolated from Taxus brevifolia (pacific yew) in 1971 (Fischer

and Ganellin, 2006) and has since become one of the most widely used chemotherapeutics to

treat a variety of common solid tumors (Hagiwara and Sunada, 2004; Speyer et al., 2017). It

induces mitotic arrest and consequently apoptosis in rapidly dividing cancer cells by binding

along the length of microtubules, thereby stabilizing them, promoting microtubule assembly, and

suppressing microtubule dynamics (Schiff et al., 1979; Kumar, 1981; Howard and Timasheff,

1988; Wang et al., 2016; Zhu et al., 2016) by strengthening the lateral protofilament interactions

between tubulin dimers (Downing and Nogales, 1998; Nogales et al., 1999). There is also

evidence that taxol decreases the number of protofilaments and increases filament flexibility

(Dye et al., 1993; Felgner et al., 1996; Díaz et al., 1998). What is crucial for the elimination of

malignant tumor cells is, however, detrimental for other, healthy cells within the body that are

also susceptible to taxol, e.g. in neurons, where it leads to microtubule hyper-stabilization and

subsequent axonal degeneration (Shichinohe et al., 2015; Tasnim et al., 2016; Gornstein and

Schwarz, 2017). Although taxol is known to cause an array of sometimes severe side-effects

(e.g. sensory and painful neuropathy, Kuroi and Shimozuma, 2004; Mielke et al., 2006;

Scripture et al., 2006; Reyes-Gibby et al., 2009; Li et al., 2015; Gornstein and Schwarz, 2017), it

has recently been suggested to exploit its microtubule-stabilizing characteristics for the

treatment of diseases other than cancer (reviewed in Baas and Ahmad, 2013),

neurodegenerative diseases (Zhang et al., 2005; Michaelis et al., 2006; Ballatore et al., 2012),

as well as nerve injury (Hellal et al., 2011; Sengottuvel et al., 2011), despite the fact that it has a

poor blood-brain-barrier permeability (Brunden et al., 2011). The microtubule-stabilizing feature

of taxol has been exploited for decades to stabilize microtubules assembled from purified tubulin

dimers in vitro (Nitzsche et al., 2010)

Microtubule gliding assays have been used in a wide range of studies for varying

purposes, including understanding the active self-assembly of the transport machinery (Lam et

al., 2016), or different nanotechnological applications (Bachand et al., 2014; Hess and Saper,

2018), such as high-throughput compound screening for modulators of motor activity (Korten et

al., 2018) or establishing molecular detection devices for e.g. diagnostic purposes (Korten et al.,

2010). A microtubule gliding motility assay thereby provides insights into the biophysical

properties of molecular motors, but, when additional proteins are introduced, also makes it

possible to investigate potential direct interactions of these proteins with microtubules and/or

motors (Howard et al., 1989; Böhm et al., 1997; Korten and Diez, 2008; Scharrel et al., 2014).



While many studies previously used gliding assays to investigating the role of MAPs like tau on

microtubule-dependent motor motility (Peck et al., 2011; Yu et al., 2014), a potential interaction

of FUS with microtubules or motors has not been evaluated in this setup to date. Microtubule

gliding assays therefore provide a suitable tool to model the translocation of motors along

microtubules, and hence to model microtubule-dependent axonal transport. In contrast to in vivo

model systems, the direct and sole interaction between proteins and microtubules or motors can

be investigated. This allows for studying the influence of either a single protein of interest (i.e.

recombinant protein) or complex solutions comprising a mixture of proteins (i.e. cell lysates) on

microtubule-dependent axonal transport. Thereby, using microtubule gliding assays,

disturbances in the axonal transport machinery can be investigated which might otherwise be

masked in in vivo models of axonal transport (e.g. cell culture models).

1.4 Aim of this study

Although axonal transport defects have long been described as a pathological hallmark of FUS-

ALS, little is known about the cause and exact underlying mechanisms. FUS protein colocalizes

with kinesin-1 mRNA and was isolated as part of an RNA-transporting granule associating with

kinesin-1 protein (Kanai et al., 2004; Yasuda et al., 2017), but whether FUS directly interacts

with kinesin-1 or microtubules remains unknown to date. This thesis hence aims at developing a

novel and robust assay mimicking axonal transport in vitro in a complex environment such as

neuronal whole cell lysates. Using this assay, two hypotheses on the pathomechanisms that

might contribute to axonal transport defects will be tested.

I) FUS directly interacts with kinesin-1 or microtubules in order to transport e.g. RNA, and its

mislocalized, mutant form directly alters this interaction and impairs kinesin-1 motility on

microtubules. This would indicate that disturbances in axonal transport are a direct

consequence of cytoplasmic mislocalization of FUS.

II) An increased ratio of 4R:3R tau isoforms, as previously observed in FUS-ALS, is sufficient to

impair kinesin-1 motility on microtubules, indicating an indirect disturbance of axonal transport

as a consequence of the nuclear loss-of-function of FUS.

To address these hypotheses, axonal transport will be modelled utilizing a

kinesin-1-dependent microtubule gliding assay. The assay will be modified to be compatible with

and provide robust results in the presence of physiological buffers and complex solutions such

as whole cell lysates. First, in order to test the direct interference of wildtype FUS-GFP or the

FUS-P525L-GFP variant with kinesin-1 and/or microtubules, recombinant FUS-GFP variants, as



well as cell lysates generated of spinal motor neurons differentiated from isogenic iPSC lines

expressing the same FUS variants, will be generated and administered to the modified gliding

assay. Second, to test the influence of increasing 4R:3R tau isoform ratios on the kinesin-1

motility on microtubules, recombinant human 2N4R tau-mScarlet and 2N3R tau-GFP isoforms

will be purified from insect cells and administered individually or combined at different ratios to

the modified kinesin-1-dependent microtubule gliding assay.

In both scenarios, the relative microtubule gliding velocity in the presence or absence of

proteins is assessed, which is determined as a measure for kinesin-1 motility. In addition, the

binding of recombinant proteins is either qualitatively assed by widefield-epifluorescence

microscopy (in the case of recombinant FUS variants) or quantitatively by total internal reflection

fluorescence (TIRF) microscopy (in the case of recombinant tau variants).

This study will hence investigate the direct or indirect interference of ALS-associated FUS

on kinesin-1-dependent transport. It will thereby provide important contributions to

understanding the pathomechanisms leading to axonal transport defects in FUS-ALS.

Mechanisms of Axonal Transport in ALS Materials and Methods


2. Materials and Methods

2.1 Materials

2.1.1 Technical equipment

Table 1: Technical equipment

Instrument Company

70 mL Polycarbonate Bottle Assembly Beckman Coulter GmbH, Krefeld, Germany

ÄKTA pure protein purification system General Electric Company, Boston, MA, USA

Analytical Balance - CP225D-0CE Analytical Balance - CP225D-0CE

Axio Observer Z1 microscope with a 1. 63x oil

immersion objective

Carl Zeiss, Jena, Germany

Azure c600 Gel Imaging System Azure Biosystems Inc., Dublin, CA, USA

Balance - SBA 52 Scaltec Instruments GmbH, Heiligenstadt,

Germany

BAT-10 Multipurpose Thermometer Physitemp Instruments, LLC, Clifton, NJ, USA

Biosafety cabinet HERAsafe® HS Heraeus Holding GmbH, Hanau, Germany

Cell culture Microscope – Axiovert 35 Carl Zeiss, Jena, Germany

Centrifuge – 5403 Eppendorf, Hamburg, Germany

Centrifuge - MiniSpinPlus Eppendorf, Hamburg, Germany

Centrifuge Heraeus® Biofuge® Pico Thermo Fisher Scientific, Massachusetts, USA

Centrifuge Heraeus® Biofuge® Primo Thermo Fisher Scientific, Massachusetts, USA

Centrifuge Heraeus™ Biofuge™ Stratos™ Thermo Fisher Scientific, Massachusetts, USA

CORIO CP-200F Thermostat JULABO GmbH, Seelbach, Germany

Elektro Automatik EA-PS 3016-10B Power

Supply

Elektro-Automatik GmbH & Co. KG, Viersen,

Germany

EmulsiFlex-C5 High Pressure Homogenizer AVESTIN Inc., Ottawa, ON, Canada

Fluorescence Microscope - Observer.Z1 Carl Zeiss, Jena, Germany

Fluorescence Microscope – Ti-E Nikon GmbH, Düsseldorf, Germany

Freezer -80°C HERAfreeze® Thermo Fisher Scientific, Massachusetts, USA

Hemocytometer - Neubauer improved Paul Marienfeld GmbH Co. KG, Lauda-

Königshofen, Germany

Hybridization oven - OV3 Biometra GmbH, Göttingen, Germany

iBlot® Gel Transfer Device Thermo Fisher Scientific, Massachusetts, USA



Imager - LAS3000 Fujifilm, Tokyo, Japan

Incubator - Heracell 150 Thermo Fisher Scientific, Massachusetts, USA

iXon EM+ DU-897 BV back-illuminated

EMCCD camera

Andor Technology Ltd., Belfast, UK

iXon Ultra back-illuminated EMCCD camera Andor Technology Ltd., Belfast, UK

Liquid Nitrogen Tank K Series Worthington Industries, Columbus, Ohio, USA

Microcentrifuge Heraeus™ Fresco™ 17 Thermo Fisher Scientific, Massachusetts, USA

Microscope CKX41 Olympus Deutschland GmbH, Hamburg,

Germany

MINIPULS 3 Peristaltic Pump Gilson Incorporated, Middleton, WI, USA

Multi-channel micropipette Transferpette®-8

electronic

Brand GmbH & Co. KG, Wertheim, GER

neoVAQ Maxi Vacuum Pump neoLab Migge GmbH, Heidelberg, Germany

Nikon Eclipse Ti microscope with a 1.49

PlanApo 100x oil immersion objective

Nikon, Tokio, Japan

OptimaTM LE-80K Ultracentrifuge

with Type45 Ti Rotor

Beckman Coulter GmbH, Krefeld, Germany

pH meter - inoLab ph 720 Wtw GmbH, Weilheim, Germany

Sunrise™ Absorbance Reader Tecan Trading AG, Switzerland

Thermomixer – Thermomixer 5436 Eppendorf, Hamburg, Germany

Thermomixer – Thermomixer Comfort Eppendorf, Hamburg, Germany

Water bath Julambo SW22 JULAMBO Labortechnik GmbH, Seelbach,

Germany

Water purification system - GenePure Thermo Fisher Scientific, Massachusetts, USA

XCell SureLock Mini-Cell Electrophoresis

System

Thermo Fisher Scientific, Massachusetts, USA

2.1.2 General equipment

Table 2: General equipment

Equipment Company

15 mL and 50 mL Falcons BD, New Jersey, USA

96-Well White Clear Flat Bottom Plates Corning Inc., Tewksbury MA, USA

Amicon® Ultra-4 centrifugal filters Ultracel® 30K Merck Millipore Ltd., Tullagreen, Ireland



Axygen® Filter Tips 10-200 µL Corning Inc., Tewksbury MA, USA

BD Luer-Lok 50 mL syringe Becton Dickinson S.A., Madrid, Spain

BD MicrolanceTM Canula 27 G 3/4 0.4 x 19 mm Becton Dickinson GmbH, Heidelberg,

Germany

Biosphere® Filter Tips 1000µl Sarstedt AG & Co, Nürnberg, Germany

Bio-Spin® Columns Bio-Rad Laboratories Inc., Hercules, CA, USA

Cell scraper TPP, Trasadingen, Switzerland

Costar® 6-well cell culture plates Corning Inc., Tewksbury MA, USA

Cryotubes Greiner Bio-One, Frickenhausen, Germany

Disposable Serological Pipettes Corning Inc., Tewksbury MA, USA

Eppendorf Tubes, 1.5mL, 2mL Eppendorf AG, Hamburg, Germany

iBlot® Nitrocellulose transfer stack Thermo Fisher Scientific, Massachusetts, USA

Mikrozid® Liquid Schülke&Mayr, Norderstedt, Germany

Millex HV low binding PVDV membrane filters

0.45 μm

Merck Millipore, Billerica, USA

Nalgene® Mr. Frosty Freezing Sigma Aldrich, St. Louis, USA

Omnifix® Syringes, 2-3 mL B. Braun Medical Inc., Bethlehem, PA, USA

Parafilm® Carl Roth GmbH + Co. KG, Karlsruhe,

Germany

ROTI®Nanoquant Protein Assays Carl Roth GmbH + Co. KG, Karlsruhe,

Germany

SuperdexTM 200 Increase 10/300 GL GE Healthcare Bio-Sciences AB, Uppsala,

Sweden

Surgical mask Paul Hartmann AG, Heidenheim, Germany

T25 Cell Culture Flasks Thermo Fisher Scientific, Massachusetts, USA

2.1.3 Cell culture ingredients, chemicals and reagents

Table 3: Cell culture ingredients, chemicals and reagents

Chemical Company

Accutase Life Technologies Corporation, Carlsbad, USA

Activin A Biomol GmbH, Hamburg, Germany

Amersham ECL Prime Western Blotting

Detection Reagent

GE Healthcare Life Sciences, Marlborough,

MA, USA



Ascorbic Acid Sigma-Aldrich, St. Louis, USA

B-27 Supplement minus Vitamin A Life Technologies Corporation, Carlsbad, USA

Benzonase(R) Nuclease Sigma-Aldrich, St. Louis, USA

BSA Fraction V Thermo Fisher Scientific, Massachusetts, USA

CHIR99021 Cayman Chemical, Michigan, USA

Chromatography paper grade 3mm GE Healthcare UK Limited, Buckinghamshire,

UK

cOmplete EDTA -free Protease Inhibitor

Cocktail Tablets

Roche Holding AG, Basel, Switzerland

DAPT Cayman chemical company, Ann Arbor, USA

dbcAMP Sigma-Aldrich, St. Louis, USA

Dichlorodimethylsilane Sigma-Aldrich, St. Louis, USA

DMEM/F-12 Life Technologies Corporation, Carlsbad, USA

DMSO Sigma-Aldrich, St. Louis, USA

DPBS without Ca2+/Mg2+ Life Technologies Corporation, Carlsbad, USA

Ethanol VWR International GmbH, Dresden, Germany

Glycerol MP biomedicals, Santa Ana, USA

Glycine Carl Roth GmbH & Co. KG, Karlsruhe,

Germany

HaltTM Phosphatase inhibitor Cocktail (100x) Thermo Fisher Scientific, Massachusetts, USA

HaltTM Protease Inhibitor Cocktail (100x) Thermo Fisher Scientific, Massachusetts, USA

HisTrap™ HP Columns GE Healthcare Bio-Sciences AB, Uppsala,

Sweden

Hydrofluoric acid ACS reagent, 48 % Sigma-Aldrich, St. Louis, USA

Imidazole Sigma-Aldrich, St. Louis, USA

Isopropanol VWR International GmbH, Darmstadt,

Germany

Kel-F Hub needle, 30 gauge, 2 inches, point

style 3

Hamilton Bonaduz AG, Bonaduz, Switzerland

KnockoutTM DMEM Thermo Fisher Scientific, Massachusetts, USA

Laminin from EHS sarcoma (mouse) Sigma-Aldrich, St. Louis, USA

Laminin from EHS sarcoma (mouse) Roche Holding AG, Basel, Switzerland

Luer-LokTM syringe Becton, Dickinson and Company, Franklin

Lakes, NJ, USA



Magnesium chloride hexahydrate Merck Millipore, Billerica, USA

MatrigelTM Basement Membrane Corning Inc., Tewksbury MA, USA

MenzelTM glass coverslips, 18 x 18 mm,

22 x 22 mm

Gerhard Menzel Glasbearbeitungswerk GmbH

& Co. KG, Braunschweig, Germany

Mikrozid Schülke&Mayr, Norderstedt, Germany

Monoclonal Anti-β-Tubulin I antibody,

clone SAP.4G5

Sigma-Aldrich, St. Louis, USA

N2-Supplement Life Technologies Corporation, Carlsbad, USA

NeurobasalTM Medium Life Technologies Corporation, Carlsbad, USA

Ni-NTA agarose beads Qiagen, Hilden, Germany

Non-Silicone Thermal Interface Compound Electrolube, Leicestershire, UK

Novex™ MagicMark™ XP Western Protein

Standard


Novex™ NuPAGE™ LDS-Probenpuffer (4X) Thermo Fisher Scientific, Massachusetts, USA

Novex™ SeeBlue™ Plus2 Prestained

Proteinmarker


Novex™ Sharp Pre-stained Protein Standard Thermo Fisher Scientific, Massachusetts, USA

NuPAGE™ MOPS SDS Running Buffer Thermo Fisher Scientific, Massachusetts, USA

NuPAGE™ Novex™ 4-12 % Bis-Tris-

Proteingel


Parafilm Bemis Company Inc., Neenah, WI, USA

Penicillin-Streptomycin Life Technologies Corporation, Carlsbad, USA

Phosphatase Inhibitor (100x) Thermo Fisher Scientific, Massachusetts, USA

Phosphate Buffer Saline (PBS) Thermo Fisher Scientific, Massachusetts, USA

Pierce™ BCA Protein Assay Kit Thermo Fisher Scientific, Massachusetts, USA

Pluronic F127 Sigma-Aldrich, St. Louis, USA

Poly-L-Ornithine Sigma-Aldrich, St. Louis, USA

Potassium chloride Merck Millipore, Billerica, USA

PreScission (HRV 3C + GST) Protein Expression Facility, MPI-CBG,

Dresden Germany

Purmorphamine Cayman Chemical, Michigan, USA

Retinoic Acid Sigma-Aldrich, St. Louis, USA

rhBDNF Promega GmbH, Mannheim, Germany

rhGDNF Sigma-Aldrich, St. Louis, USA



SAG Cayman chemical company, Ann Arbor, USA

SDS Sigma-Aldrich, St. Louis, USA

Sf9-ESF Expression Systems, LLC, Davis, CA, USA

SimpleBlue Safe Stain Invitrogen, Carlsbad, CA, USA

SuperSignal™ West Femto Maximum

Sensitivity Substrate


TGFβ-3 Peprotech, London, UK

Tris-HCL Carl Roth GmbH & Co. KG, Karlsruhe,

Germany

Tween 20 Serva Elektrophoresis GmbH, Heidelberg,

Germany

β-Mercaptoethanol Sigma-Aldrich, St. Louis, USA

2.1.4 Media and buffers

Table 4: Media and buffers, all amounts given in percent refer to volume/volume

Medium Composition

N2B27 medium 48.75 % DMEM/F12, 48.75 % NeurobasalTM Medium, 1 %

Penicillin/Streptomycin, 1 % B-27 Supplement without

Vitamin A, 0.5 % N2-Supplement

Normal culture (expansion)

medium

N2B27 medium with 3 µM Chiron99021, 150 µM AA,

0.5 µM SAG

Patterning medium N2B27 medium with 1 ng/mL rhBDNF, 0.2 mM AA, 1 µM

retinoic acid, 1 ng/mL rhGDNF, 0.5 µM SAG

Maturation medium N2B27 medium with 0.1 mM cAMP, 2 ng/mL rhBDNF,

0.2 mM AA, 1 ng/mL TGFβ-3, 2 ng/mL rhGDNF, 5 ng/mL

Activin A (on day 1 of maturation), 5 µM DAPT (on day 4

of maturation)

Lysis buffer for whole cell lysates PBS with 1 x protease inhibitor, 10 % glycerol, 10 mM β-

glycerophosphate

FUS-GFP storage buffer 50 mM Tris-HCL, 500 mM KCL, 1 mM DTT, 5 % glycerol

in H2O

BRB80 80 mM PIPES, 1 mM EGTA, 1 mM MgCl2

Standard motility buffer PBS with 10 µM taxol, 10 % glycerol, 10 mM β-

glycerophosphate, 0.3 % methylcellulose, 1 mM ATP,



20 mM D-glucose, 20 μg/mL glucose oxidase, 10 μg/mL

catalase, 10 mM DTT (and 0.2 mg/mL casein in absence

of recombinant protein, BSA, or cell lysate)

VALAP Vaseline, lanolin, and paraffin in a 1:1:1 ratio

Transfer buffer for western blot 200 mM glycine, 25 mM Tris-HCL, 6.7 % methanol (in

cathode buffer) or 0.1 % SDS (in anode buffer) in H2O, pH

8.3

PBST PBS with 0.05 % Tween20

Stripping buffer 3 g glycine, 0.1 % SDS, 1 % Tween20, in H2O, pH 2.2

HisTrap storage buffer 2x 50 mM HEPES, 300 mM KCL, 0.2 % Tween20 (v/v) in

H2O

Equilibration buffer

HisTrap storage buffer with 30 mM imidazole, 10 mM β-

mercaptoethanol in H2O

Lysis buffer for the purification of

tau-GFP from Sf9 cells

50 mL equilibration buffer with one protease inhibitor

tablet

High salt buffer Equilibration buffer with 200 mM KCL

Elution buffer Equilibration buffer with 200 mM imidazole

2.1.5 Antibodies

Table 5: Primary antibodies

Antibody Host Dilution Company Reference Number

FUS Mouse 1:1000 Sigma-Aldrich, St. Louis, USA AMAB90549-100UL 3R Tau Mouse 1:2000 Merck KGaA, Darmstadt,

Germany 05-803

4R Tau Mouse 1:2000 Merck KGaA, Darmstadt, Germany

05-804

β-actin Rabbit 1:8000 Cell Signaling Technology, Inc, Beverly, MA, USA

#4967

Table 6: Secondary antibodies

Antibody Host & Reactive Species

Dilution Company Reference Number

HRP anti Mouse Donkey 1:6000 DIANOVA GmbH, Hamburg, Germany

711-035-150

HRP anti Rabbit Donkey 1:6000 DIANOVA GmbH, Hamburg, Germany

711-035-152



2.1.6 Software

Table 7: Software

Product Type Company

FIJI (ImageJ) 1.52p Image analysis Open source (Schindelin et al., 2012)

Matlab R2017a Data evaluation and graphics The MathWorks, Inc., Natick, MA,USA

Inkscape 0.91 Figure preparation Open source (www.inkscape.org)

GraphPad Prism 7 Statistical analysis GraphPad Software Inc., La Jolla, USA

Microsoft Office 2010 Data analysis and writing Microsoft Corporation, Redmond,

Washington, USA

MetaMorph 7.8.4.0 Image analysis Molecular Devices, Sunnyvale, CA,

USA

Image StudioTM Lite,

5.2.5

Western blot image analysis LI-COR Biosciences, Lincoln, Nebraska

USA

OriginLab 2019b Statistical analysis OriginLab Corporation, Northampton,

MA, USA

2.1.7 Patient-derived isogenic iPS cell lines

Isogenic cell lines used in this study were generated and characterized previously by members

of the group of Prof. Andreas Hermann. The specifications for both lines are summarized in

Table 8. This study was approved by the local ethics committee (EK45022009, EK393122012).

Table 8: Characteristics for isogenic cell lines used in this study

Name Genotype Gender Year of

birth Age at biopsy Reference

Wildtype

FUS-EGFP

FUS-WT-

EGFP/WT F 1952 58 (Naumann et al., 2018)

FUS-P525L-

EGFP

FUS-P525L-

EGFP/WT F 1952 58 (Naumann et al., 2018)



2.2 Methods

2.2.1 Coating of cell culture plates and flasks

NPCs were maintained and primed for differentiation in T25 flasks. Flasks were coated with

Matrigel diluted 1:100 in KnockOut™ DMEM and incubated at room temperature (RT) for

30 min. For maturation, cells were cultured on 6-well plates coated first with 15 % Poly-L-

Ornithine (PLO) in PBS, then with 0.005 mg/ml laminin in PBS at 37 °C overnight, respectively.

2.2.2 NPC thawing, culture, and cryoconservation

NPCs were thawed by suspending them in pre-warmed N2B27 medium, centrifuged with

230 x g for 5 min and subsequent seeded into Matrigel-coated culture flasks in expansion

medium. Cells were maintained at standard cell culture conditions (37 °C in a humidified

atmosphere containing 5 % CO2) in N2B27 medium supplemented with 3 µM Chiron99021,

150 µM ascorbic acid (AA), and 0.5 µM smoothened agonist (SAG). Medium was changed three

times a week, while cells were split at 85 % confluency, on average once per week. For splitting,

cells were detached from the surface via incubation with accutase at 37 °C for 5 min. Detached

cells were taken up in pre-warmed N2B27 medium and either reseeded at an appropriate

density or cryopreserved at -80°C. For this, cells were resuspended in N2B27 medium

containing 10 % DMSO as a cryoprotectant. Cells were frozen over night at a cooling rate of

~1 °C per hour, and subsequently transferred to liquid nitrogen (-180 °C) for long-term storage.

2.2.3 Differentiation and maturation of NPCs towards spinal motor neurons

The generation of spinal motor neurons out of NPCs requires the addition of different small

molecules to the culture medium at specified time points (Figure 5). After splitting, NPCs were

reseeded in N2B27 medium supplemented with 1 ng/mL recombinant human brain-derived

neurotrophic factor (rhBDNF), 0.2 mM AA, 1 µM retinoic acid, 1 ng/mL recombinant human

glial-derived neurotrophic factor (rhGDNF), and 0.5 µM SAG in order to prime their

differentiation into spinal motor neurons. Medium was exchanged every two days until day six

post induction, where the maturation phase was started by the addition of N2B27 medium

supplemented with 0.1 mM cAMP, 2 ng/mL rhBDNF, 0.2 mM AA, 1 ng/mL Transforming growth

factor β-3 (TGFβ-3), and 2 ng/mL rhGDNF, which marked day 0 of maturation. On this day only,

5 ng/mL Activin A was additionally supplemented to the maturation medium. Primed NPCs were

split one more time between days 2-4 of maturation and reseeded in maturation medium into a

6-well plate coated with PLO and laminin, at a concentration of 1 x 106 cells/well (determined

using a Neubauer counting chamber). On day 4 of maturation only, 5 µM N-[N-(3,5-



Difluorphenacetyl)-L-alanyl]-S-phenylglycin-tert-butylester (DAPT) was added to the maturation

medium. DAPT is a γ-secretase inhibitor that inactivates Notch signaling and thereby results in

fewer cells in culture with progenitor morphology while increasing cells with a neuronal

morphology (Nelson et al., 2007). Maturing spinal motor neurons were kept in maturation

medium without any additional supplements, with medium changes twice a week, until they

were lysed on day 21 of maturation.

Figure 5: Timeline for the differentiation and maturation of NPCs towards spinal motor neurons

Different sets of small molecules are added to N2B27 medium during the differentiation and maturation of NPCs towards spinal motor neurons.

2.2.4 Generation of cell lysates – isolation techniques

At day 21 of maturation, spinal motor neurons were gently washed with PBS once, scrapped off

the dish surface and resuspended in fresh PBS. Cells were then lysed by following methods:

A | Chemical lysis using the Active Motif® Nuclear Extraction Kit according to the

manufacturer’s instructions for the generation of whole cell lysates. For the exchange of buffers

after chemical lysis, PD Minitrap™ G-10 desalting columns or Vivaspin® 500 centrifugal

concentrators were used according to the manufacturer’s instructions. Protein in PD Minitrap

columns was eluted in PBS. Centrifugation speed and time for Vivaspin columns was 13.000 x g

and 30 min, respectively.

B | Lysis by grinding in liquid nitrogen: Cells were pelleted at 14.000 x g for 5 min at 4°C

and the cell pellet was resuspended in 1 ml fresh PBS supplemented with 1 x protease inhibitor

cocktail (containing 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride, aprotinin, bestatin,

E-64, leupeptin and pepstatin A), dropwise frozen in liquid nitrogen and ground to a fine powder

using a pre-chilled mortar.



C | Mechanical lysis with glass beads: Cells were pelleted at 14.000 x g for 5 min and the

cell pellet resuspended in 1 ml fresh PBS supplemented with 1 x protease inhibitor. Sterile glass

beads were added to each Eppendorf tube to an approximate volume of 400 µL, and the tube

was vortexed for 5 min. Glass beads were allowed to sediment on ice for 5 min and the

supernatant was aspirated.

D | Mechanical lysis using a dounce homogenizer: Cells were pelleted at 14.000 x g for

5 min and the cell pellet was resuspended in 1 ml fresh PBS supplemented with 1 x protease

inhibitor. Resuspended cell were sheared in a dounce homogenizer on ice using ten strokes.

E | Mechanical lysis using a syringe: Cells were pelleted at 14.000 x g for 5 min. The cell

pellet was resuspended in 1 ml fresh PBS supplemented with 1 x protease inhibitor and cells

were sheared by pulling them ten times through a thin needle (0.4 x 19 mm; 27G ¾) using a

2 ml syringe while on ice.

Lysates prepared by methods B through E were subsequently centrifuged at 4 °C and

70.000 x g for 5 min. The supernatant was then transferred to a new tube, supplemented with

1 x protease inhibitor, 10 mM β-glycerophosphate, and 10 % glycerol, and snap-frozen in liquid

nitrogen for long term storage at -80 °C.

2.2.5 Determining protein concentration

Protein concentrations were determined using the Pierce™ bicinchoninic acid (BCA) protein

assay kit (Thermo Fisher Scientific) according to the manufacturer’s instructions. Briefly,

standards were prepared with BSA diluted in lysis buffer to concentrations ranging from

25 µg / ml to 1 mg / m and measured in duplicates in a white 96-well transparent flat bottom

plate. Lysates were measured in triplicates. All probes were mixed with the kit’s working reagent

within the plate and subsequently incubated at 37 °C for 30 min. After incubation, the plate was

allowed to cool down to RT, the absorbance of samples at 562 nm was measured with a

TECAN plate reader and corrected for the values of blank measurements. A standard curve was

created in Microsoft Excel 2010, and the sample concentration was calculated from a linear

regression line. The BCA assay is a highly sensitive, two-step reaction. In the first step, cupric

ions (Cu2+) are reduced to cuprous ions (Cu+) by proteins under alkaline conditions, a reaction

referred to as the Biuret reaction (Gornall et al., 1949), while two BCA molecules are chelated in

the second step by one cuprous ion, resulting in a water-soluble, stable BCA/copper complex

(Smith et al., 1985). This complex absorbs light at 562 nm, which causes a color change of the

solution from blue to intense purple, measurable using a spectrophotometer. The measured



absorbance is directly proportional to the amount of protein in solution. The assay allows for the

measurement of protein over a wide range of concentrations (between 0.5 ng/µL and 1.5 µg/µL)

and is compatible with many detergents and other substances. However, exogenous reducing

agents, such as DTT or mercaptoethanol, or copper chelating agents in particular greatly

interfere with the reaction and hence may influence the results. Similarly, the presence of

certain amino acids (i.e. cysteine, tyrosine, and tryptophan, Wiechelman et al., 1988) influences

the accuracy of the results. In addition, the assay requires comparably long incubation times of

30 min up to two hours and incubation at 37°C, as the reaction is temperature sensitive (Lowry

et al., 1951; Smith et al., 1985; Krohn, 2002; Brady and Macnaughtan, 2015).

For reasons of comparison, the protein concentration of lysates produced by chemical

lysis was additionally determined using the ROTI®Nanoquant Bradford assay in a microtiter

plate according to the manufacturer’s instructions. In brief, standards were prepared with BSA

diluted in PBS ranging from a concentration of 1 to 100 µg / ml and again prepared in duplicates

in a white 96-well transparent flat-bottom plate. Lysates were diluted 1:20 and measured in

triplicates. All probes were then mixed with working reagent diluted 1:5 in H2O and incubated at

RT for 5 min, after which absorbance was measured three times at 450 nm and 590 nm, each

and respectively, in a TECAN plate reader. For each well, the measured value at 590 nm was

divided by the value measured at 490 nm, and the average of three subsequent measurements

was taken to calculate the final concentration. A standard curve was created in this way,

through which the sample concentration could be estimated using a linear regression line. The

Bradford assay is comparably sensitive, but more straightforward as it deploys the ability of the

dye Coomassie blue to directly bind to proteins and subsequently change its color from red

(absorbance at 465 nm) to blue (absorbance at 595 nm). The absorbance can be measured at

both wavelengths, and the change in color is directly related to the amount of protein present in

the sample. Since this method does not require a series of reactions to indicate protein

concentration, the incubation time is much faster (5 min) compared to BCA, although the range

of protein amount that can be detected is limited (between 1 and 20 µg). The reagent is

however compatible with a large variety of buffers and solvents, as well as reducing and

chelating agents, which makes it a suitable alternative to BCA if such agents cannot be

removed from solution. The downside is that the Coomassie reagent precipitates in the

presence of surfactants and is highly acidic, indicating that proteins with poor acid solubility

cannot be measured with this dye. Similar to BCA, it is also very sensitive to certain amino acids

(i.e. aromatic acids such as arginine, histidine or lysine, Compton and Jones, 1985; de Moreno

et al., 1986; Fountoulakis et al., 1992). In addition, it adsorbs to glassware or quartz cuvettes,



which require thorough cleansing after detection in order to be reusable (Bradford, 1976; Zor

and Selinger, 1996; Krohn, 2002; Georgiou et al., 2008; Brady and Macnaughtan, 2015).

2.2.6 Acquisition of kinesin-1, tubulin, and recombinant human FUS-GFP variants

Porcine tubulin was purified from porcine brain (obtained from Vorwerk Podemus, Dresden,

Germany) using previously established protocols (Castoldi and Popov, 2003) and fluorescently

labeled using mixed isomers of 5-(and 6-) carboxytetramethylrhodamine, succinimidyl ester

(5(6)-TAMRA,SE, Invitrogen, C1171), a member of the rhodamine family of dyes (Gessner and

Mayer, 2000). Drosophila melanogaster kinesin-1 heavy chains were expressed in full length in

insect cells and purified as previously described (Korten et al., 2016). Recombinant human

FUS-GFP variants were kindly provided by Dr. Jie Wang, Max Planck Institute of Molecular Cell

Biology and Genetics, Dresden. The first batch of recombinant human 2N4R tau-GFP protein

used for the determination of assay sensitivity was kindly provided by Dr. Marcus Braun,

Institute of Biotechnology CAS, Prague.

2.2.7 Expression and purification of recombinant human tau variants

The construct for recombinant human 2N3R tau-GFP was kindly provided by Prof. Eckhard

Mandelkow, Deutsches Zentrum für Neurodegenerative Erkrankungen e.V., Bonn and is

described in Gustke et al., 1994 and Seitz et al., 2002. The construct for recombinant human

2N4R tau-GFP was kindly provided by Dr. Amayra Hernandez Vega, Max Planck Institute of

Molecular Cell Biology and Genetics, Dresden, and is characterized in Hernández-Vega et al.,

2017. In order to express both constructs in insect cells, the gene of interest was amplified by

PCR and flanked with the restriction sites NotI and AscI using the respective

5’ (AATAATAACATGCGGCCGCAATGGCTGAGCCCCGCC) and

3’ (AATAATAACATGGCGCGCCCAAACCCTGCTTGGCCAG) primers. The resulting product

was inserted into the vector pOCC61, containing a C-terminal monoGFP tag and a

3C-cleavable 6xHis-tag for affinity purification. Baculovirus for the expression in insect cells was

generated from these constructs by Régis Lemaitre at the Protein Expression Purification and

Characterization (PEPC) facility at the Max Planck Institute of Molecular Cell Biology and

Genetics, Dresden (Lemaitre et al., 2019). Insect cells (Sf9-ESF) were infected with the

respective virus with a ratio of 1:100 (v/v) and harvested 72 hours post infection by

centrifugation at 350 x g for 10 min.



Purification of recombinant human tau-GFP variants

Harvested cell pellets were resuspended in 20 mL lysis buffer (50 mL equilibration buffer with

one protease inhibitor tablet) and the suspension lysed by passing them three times through an

Emulsiflux French press on ice. All further steps were performed at 4°C or on ice. The lysate

was collected in cold Beckmann centrifuge bottles, supplemented with benzonase (at least

1:6000), and cleared of cell debris by centrifugation at 14000 x g at 4°C for 45 min. The cleared

lysate was then passed through a 0.45 µm filter in order to completely remove cell debris before

protein purification via nickel-sepharose affinity chromatography. His-trap columns were

equilibrated with 10 column volumes (CV) of equilibration buffer before the protein was loaded

onto the column by passing the supernatant through the column at a flow rate of about

0.75 ml/min with the help of a peristaltic pump. The column was washed once with 10 CV high

salt buffer or equilibration buffer (50 mM HEPES, 300 mM KCL, 0.2 % Tween20 (v/v),

10 mM β-mercaptoethanol in H2O) without imidazole. The protein was then eluted by flowing

elution buffer through the column and collecting 1 ml fractions. Eluted fractions were pooled and

supplemented with PreScission 3C-6xHis protease (1 mg/ml) for GST(His)-tag removal, shaking

at 4°C overnight. On the next day, the sample was concentrated and the elution buffer

exchanged for equilibration buffer by centrifugation through 30K AmiconUltra filters at 7000 rpm

for 10 min three times. In order to remove the cleaved His-tag from the cleaved protein fraction,

the concentrated sample was incubated with 1 mL Ni-NTA agarose beads shaking at 4°C for

one hour. This solution was then passed through a 10 mL gravity flow filter column which

retained the beads and the flow through was concentrated again by centrifugation as described

above. 50 µL samples were collected after each step to determine purification efficiency in a

subsequent sodium dodecyl sulfate-polyacrylamide (SDS-PAGE) gel (Figure S1). The protein of

interest was further purified by size exclusion chromatography (SEC) with equilibration buffer

through a superdex200 column on an ÄKTA pure liquid chromatography system at 4°C.

Absorbance was measured at 480 nM for eGFP construct and 569 nM for the mScarlet

construct and the fractions under the peak (0.5 mL per fraction) were collected. These pooled

fractions were again concentrated by centrifugation as described above, and the concentrated

protein was flash frozen in liquid nitrogen and stored at -80°C in 5 µL aliquots.

Protein quantification by SDS-PAGE gel electrophoresis and Coomassie staining

Purified proteins and samples taken throughout the purification procedure were analyzed by

SDS-PAGE for purification efficiency and concentration measurement. SDS loading buffer,

supplemented with 0.1 M DTT, was mixed 1:4 with the respective samples to be analyzed and



boiled at 95°C for 5 min. 10 µL of sample was loaded onto a 4-12 % Bis-Tris gel, together with

an appropriate protein ladder. The gel was assembled into an SDS gel electrophoresis running

chamber filled with 1 x MOPS buffer and run at 120 V, 85W, and 60 mA until all protein had

passed to the end of the gel (at least one hour). Gels were then stained with Coomassie

SimplyBlue SafeStain for at least 45 min up to overnight. Afterwards, gels were destained in

ddH2O until bands were clearly visible.

Protein quantification on SDS-PAGE gels

Purified 6xHis-eGFP protein at a known concentration was run as a standard on SDS-PAGE

gels, together with different dilutions of the protein of interest and stained with Coomassie blue

as described above. Gels were scanned with an Azure c600 Gel Imaging System and the

integrated density of protein bands was quantified using ImageJ. A rectangular area was

selected enclosing the first lane. The same rectangular shape was the applied onto all lanes on

the gel. The intensity profiles of all lanes were then plotted using the ‘analyze gel’ function of

ImageJ and the integrated intensities were recorded by selecting the area under each peak. A

calibration curve was then obtained from the linear fit to the integrated intensity for different

concentrations of the 6xHis-eGFP protein. Protein sample concentration were then calculated

for each dilution and averaged to get the undiluted protein concentration.

2.2.7 Kinesin-1-dependent microtubule gliding assay

Flow chamber and imaging assembly

Kinesin-1-dependent microtubule gliding assays were adapted from previously described

protocols (Nitzsche et al., 2010). Assays were performed in manually assembled flow chambers

(Figure 6), which were constructed by cutting out seven stripes of parafilm of about 25 x 1 mm2

size and sandwiching them in between 18 x 18 mm2 (top) and 22 x 22 mm2 (bottom) glass

coverslips, forming parallel channels. In addition, two 25 x 3 mm2 parafilm stripes were cut and

placed orthogonal to the channels on the sides of the top coverslip, just above the entry and exit

sites of channels, to prevent immersion oil from the objective from flowing into the channels

during later imaging. Overhanging parafilm was cut off using a razor blade. The sandwich was

heated until the parafilm got soft and six channels were sealed leak-proof by gently applying

pressure onto the coverslips. Molecular motor motility is very sensitive to changes in

temperature (Böhm et al., 2000; Hong et al., 2016). In order to minimize temperature

fluctuations, and therefore improve reproducibility of motility measurements, an objective heater

was used at the microscope and flow chambers were mounted onto a custom-build Peltier

element (Figure 6), and both devices were set to 30 °C. The Peltier element makes use of the



Peltier effect, which describes the transfer of heat from one junction to another if a voltage is

applied across joined conductors, creating an electrical current, with consumption of electrical

energy. The effect was discovered in 1834 by Jean Charles Athanase Peltier (Peltier, 1834) and

allows for heating as well as cooling, depending on the direction of the current (Taylor and

Solbrekken, 2008). A Peltier element therefore has two major advantages: i) it does not

overheat, but can actively cool down its surface by changing the electrical polarity when

connected to a conventional power supply, and ii) reaches desired temperatures on a very fast

time scale (e.g. a temperature change of 8 °C is achieved in about 25 s) (Ionov et al., 2006;

Nitzsche et al., 2010). The temperature of the herein used Peltier element was constantly

monitored using a thermometer with analogue readout capability and the current was adjusted if

needed to maintain a steady temperature of 30 °C within flow chambers with an accuracy of

0.1 °C. The Peltier element and the flow chambers were coupled by applying, electrolube, a

silicon-free heat transfer compound, in a thin layer on the surface of the Peltier element. The

flow chamber was stuck with its bottom coverslip onto the lube and firmly attached by gentle

pressure on the flow chamber. Because of this setup, flow channels were images upside down

through the 18 x 18 mm2 top glass slip. To prevent light reflecting from the surface of the Peltier

element, bottom coverslips of all flow chambers were darkened with a black permanent marker

before fixation to the Peltier element.



Figure 6: Assembly, mounting, and imaging of flow channels.

Flow chambers are assembled by stacking two coverslips of different sizes on top of each other, with parallel stripes of parafilm in between to form individual channels. Two wider stripes of parafilm are mounted orthogonal to flow channels on the edges of the top coverslip to prevent immersion oil from flowing into the channels during imaging. By heating this assembly, the parafilm melts and tightly seals channels. The backside of the bottom coverslip was painted with a black permanent marker to prevent reflection of light after the assembly was mounted on a custom-built Peltier element, using electrolube as a temperature conducting adhesive. The Peltier element was equipped with a temperature sensor at its surface to constantly monitor the temperature. All solutions used in the experiments are pipetted onto one side of the flow chamber and aspirated at the other end of the channel using chromatography filter paper. Flow chambers in this setup were installed at the microscope upside-down, enabling imaging through the top coverslip.

Microtubule assembly and kinesin-1-dependent gliding motility assay

Microtubules were polymerized at 37 °C for 15 min from 32 μM rhodamine-labeled tubulin in

BRB80 supplemented with 4 mM MgCl2, 1 mM Mg-GTP, and 5 % DMSO in a total volume of

6.25 μL. Polymerized microtubules were immediately stabilized with taxol by adding 10 µM taxol

in BRB80 to a final volume of 100 μL, and further kept at RT in the dark. Directly before use,

microtubules were diluted 1:10 in standard motility buffer and carefully sheared twice through a

Kel-F Hub needle (gauge 30, point style 3) using a 1 mL Luer-LokTM syringe.

For the assay itself, a series of solutions was flushed through each flow channel by

carefully pipetting 20 μL of each solution on one end of the channel while sucking the previous

solution out of the channel at the other end using chromatography paper. All solutions were

applied at RT. In a first step, flow channels were perfused with a casein-containing solution



(0.5 mg/mL) in PBS and left to adsorb for 5 min in order to block unspecific protein binding sites

on the glass surface. Next, a solution containing Drosophila full-length kinesin-1 heavy chains

(4 μg/mL), casein (0.2 mg/mL), 1 mM ATP, and 10 mM DTT in PBS was flushed through the

channels and again left to adsorb for 5 min in order for kinesin-1 to attach to the glass surface.

Following this, diluted and sheared microtubules in standard motility buffer were perfused into

the channels and allowed to attach to motors for 5 min. Flow channels were then mounted onto

a Nikon Eclipse Ti microscope equipped with a Perfect Focus System (PFS) and a 1.49

PlanApo 100 x oil immersion objective heated to 30 °C, the same temperature as the Peltier

element. During imaging, the Peltier element was continuously monitored and adjusted to

ensure a constant temperature of 30 °C with a precision of 0.1 °C. Images were acquired using

widefield-epifluorescence and excited by LED lamp and respective filter sets for excitation at

550 nm for rhodamine-labeled microtubules, 470 nm for eGFP-labeled proteins, or 640 nm for

mScarlet- and mCherry-labeled proteins, respectively. Time lapse images were recorded at ten

different positions within a channel, at a rate of one frame per second and 10 frames at each

position. Images were recorded with an exposure time of 100 ms using an iXon Ultra back-

illuminated EMCCD camera in conjunction with Nikon NIS-Elements imaging software.

After imaging rhodamine-labeled microtubules alone, channels were perfused depending

on the experiment with either recombinant human tau isoforms, recombinant human wildtype

FUS-GFP or FUS-P525L-GFP, standard motility buffer, BSA, or the respective cell lysate. Each

of these solutions was supplemented with 0.3 % methylcellulose, 20 mM D-glucose, 20 μg/mL

glucose oxidase, 10 μg/mL catalase, 10 mM DTT, and 1 mM ATP to ensure optimal conditions

for efficient kinesin-1 motility. Each channel was then sealed off with VALAP (Vaseline, lanolin,

and paraffin in a 1:1:1 ratio) to inhibit evaporation. Flow chambers were allowed to equilibrate

for 5 min before running the imaging workflow as described above. In channels where tau or

FUS-GFP variants were administered, single frames were captured via excitation with 470 nm

or 640 nm light and an exposure time of 100 ms at every position immediately before acquiring

time laps movies of rhodamine-labelled microtubules. For the first batch of 2N4R tau-GFP,

single frames of the first imaged positions were captured via excitation with 470 nm light and an

exposure time of 100 ms immediately before acquiring time laps movies of rhodamine-labelled

microtubules, while single frames of the remaining nine imaging positions were captured via

excitation with 470 nm light after time lapse acquisition. This workflow takes into consideration

that analyzing the colocalization of moving microtubules with potentially bound proteins was

only possible if overlaid images were captured virtually simultaneously in both channels. In case

of FUS-GFP variants, the buffer control experiments were performed with standard motility



buffer to which FUS-protein storage buffer was added in equal dilutions as required for the

respective protein concentrations tested. When flow chambers were imaged over the course of

three hours, channels were sealed with VALAP to prevent evaporation.

2.2.8 Detection of protein binding to microtubules

Microtubules were polymerized at 37 °C for 30 min from 40 μM Atto647-labeled tubulin in

BRB80 supplemented with 4 mM MgCl2, 1 mM Mg-GTP, and 5 % DMSO in a total volume of

6.25 μL. Polymerized microtubules were immediately stabilized with taxol by adding 10 µM taxol

in BRB80 to a final volume of 100 μL, and further kept at RT in the dark. Microtubules were

cleared from free tubulin by centrifugation in a table top centrifuge at 13.3 rpm for 30 min at RT,

the supernatant was removed and the pellet resuspended in 400 μL BRB80 supplemented with

10 μM Taxol.

For the assay itself, flow chambers were assembled as described above, except that the

coverslips used for this assay were silanized with dichlorodimethylsilane (DDS) in order to

increase the hydrophobicity of the surface.

The first solution containing α-tubulin antibodies, 1:200 diluted in BRB80, had to be

sucked through each channel with the help of a vacuum pump due to the strong hydrophobicity

of the glass surface, and was left to adsorb for 15 min. Next, a solution containing the 1 % of the

non-ionic detergent F127 was flushed through the channels and again left to adsorb for at least

one hour in order to reduce non-specific protein binding to the glass surface. Flow chambers

containing F127 were then either used directly or kept in a sealed humidified container at 4°C

for up to two days. Channels were washed once with BRB80 directly before use, and then

perfused with 20 μL undiluted Atto-647-labeled microtubules for 1 min. Unbound microtubules

were then washed out with standard motility buffer without ATP, before channels were perfused

with recombinant 2N3R tau-GFP or 2N4R tau-mScarlet or standard motility buffer without ATP.

Each channel was then sealed off with VALAP to inhibit evaporation. Flow channels were then

mounted onto an Axio Observer Z1microscope equipped with a Zeiss TIRF slider and a 1.46

PlanApochromat 63 x oil immersion objective combined with a 1.6x optovar and heated to

30 °C. Images were acquired by TIRF using a 532 nm laser line for Atto647-labeled

microtubules, 488 nm for eGFP-labeled proteins, or 642 nm for mScarlet- labeled tau isoforms,

respectively, with an exposure time of 100 ms. Single images at each wavelength were

recorded at ten different positions within one channel using an iXon Ultra back-illuminated

EMCCD camera in conjunction with MetaMorph MicroManager imaging software.



2.2.9 Data analysis and statistics

Kinesin-1-dependent microtubule gliding velocities were derived from time lapse movies using

an automated MATLAB script based on a microtubule tip-tracking algorithm developed in-house

(Ruhnow et al., 2011). In detail, the position of a microtubule tip was localized in each frame

with sub-pixel resolution and compared to its position in the subsequent frame one second later,

thereby determining the frame-to-frame velocities for this microtubule between two frames. All

microtubule frame-to-frame velocities determined in each of the ten time lapse movies for one

experimental condition are combined in a MATLAB cell format for further analysis. The median

microtubule gliding velocity was determined from this data set, as well as fraction of gliding

microtubules. The median gliding velocity of microtubules in one channel after perfusion with

recombinant protein, BSA, cell lysate, or standard motility buffer, respectively, was then

normalized to the median gliding velocity of microtubules in the same channel in standard

motility buffer before perfusion. This way of analysis yields the relative speed for microtubule

gliding in a certain experimental condition, a more robust parameter for comparison among

experiments than the absolute microtubule gliding velocity. The fraction of non-gliding

microtubules describes the number of microtubules gliding with a frame-to-frame velocity less

than 20 % of the gliding velocity in standard motility buffer in that channel.

The binding affinity of 2N3R tau-GFP and 2N4R tau-mScarlet to microtubules was

determined by measuring the integrated intensity in the respective image channel along at least

20 microtubules in each of the ten positions imaged using the line tool of ImageJ, and the

fluorescence intensity along each microtubule was background corrected locally. Therefore, at

least 200 microtubules were measured per condition and experiment to obtain the average

fluorescence intensity for every tested concentration of tau isoforms. Intensities for each tau

isoform were normalized in every experiment to the respective highest concentration tested in

order to determine the fraction of bound tau protein to microtubules. The fraction of bound tau

was plotted against the concentration of tau protein in solution and fit with the Hill model using

OriginLab 2019.

Significance was determined using either one-way or two-way ANOVA and Tukey’s

multiple comparison post-hoc test (see supplementary tables). Kymographs were created using

the Kymograph plugin for ImageJ created by J. Rietdorf (FMI Basel) and A. Seitz (EMBL

Heidelberg), while overlay images were generated using MetaMorph.



2.2.10 Western blot analysis

SDS-PAGE gel electrophoresis and blotting

Probes for Western blot analysis were mixed with NuPage loading buffer in a 3:4 ratio, heated

to 95 °C for 5 min to denature all proteins and put on ice immediately after. 12 µL of this mixture

were loaded on a 4-12 % Bis-Tris SDS-PAGE gel, as well as at least one lane with 5 µL of a 1:5

mixture of two markers, one visible by eye (SeeBlue) and one visible during detection

(MagicMarker). The gel was assembled into an electrophoresis running chamber filled with

1 x MOPS buffer and electrophoresis run for at least one hour with the following parameters:

140 V, 85W, and 60 mA for one and 140 mA for two gels simultaneously. Following

electrophoresis, gels were kept in transfer buffer supplemented with 0.1 % SDS for 8 min to

increase protein release from the gel during blotting. In contrast, the PVDF membrane used to

blot on and the thick filter paper within the iBlot transfer stack were soaked in transfer buffer

containing 6.7 % methanol prior to blotting in order to activate the membrane and increase

retention of proteins on the membrane. iBlot transfer stacks were assembled according to the

manufacturer’s instructions in the iBlot transfer system and blotted for 7 min with 19 V. After

blotting, membranes were transferred into 50 mL falcons and blocked for one hour at RT with

PBST containing 5 % milk powder. Membranes were then incubated with primary antibodies

over night at 4 °C and washed twice with PBST. After incubation with horseradish peroxidase

(HRP)-coupled secondary antibodies for one hour at RT, membranes were washed again with

PBST five times before detection.

As a means to accurately determine whether the expression level of a protein of interest

differs between two samples, the detected intensity for bands for this protein of interest should

be normalized to the intensity of a band for a housekeeping protein within the same sample. In

order to not mask any bands of the protein of interest, membranes were incubated with

antibodies to detect the protein of interest, as described above, imaged, and subsequently

stripped. Membranes were stripped at RT by incubating them in fresh stripping buffer twice for

10 min each, two subsequent washing steps with PBS for 10 min each, and two final washing

steps with PBST for 5 min each. After this procedure membranes were again blocked and

incubated with respective primary and secondary antibodies for the detection of the

housekeeping protein GAPDH.

Detection, imaging and quantification of protein bands

Chemiluminescent detection was achieved by applying either Amersham ECL Prime or

SuperSignalTM West Femto substrate according to the manufacturer’s instructions, depending



on the strength of the signal as the latter is more sensitive. Imaging was immediately performed

after substrate application on an LAS3000 imager with continuous exposure and image

acquisition every 10 sec. The integrated intensity of protein bands was determined using Image

StudioTM Lite as follows: a rectangular area was created around the largest band and all other

bands subsequently selected with the same rectangular area. The background was computed

as the median pixel intensity within a three pixel area bordering the top and bottom of each

rectangular area around the corresponding band. The background value was hence calculated

for every band individually and locally and subtracted from the total intensity in the respective

rectangular area. To determine the concentration of FUS variants and tau isoforms in whole cell

lysates, recombinant FUS-WT-GFP, 2N3R tau-GFP or 2N4R tau-mScarlet of known

concentrations were loaded on SDS-PAGE gels in combination with whole cell lysates. A

standard curve was created in Microsoft Excel 2010, and the protein concentration in whole cell

lysates was calculated from a linear regression line.

Mechanisms of Axonal Transport in ALS Results


3. Results

3.1 Modification of a kinesin-1-dependent microtubule gliding assay for the

application of whole cell lysates

Defects in axonal transport have long been recognized as part of the pathology in FUS-ALS, but

only little is known about the causes and mechanisms involved in its development (De Vos and

Hafezparast, 2017a). This thesis aimed at investigating possible direct engagement of FUS

variants with the transport machinery, as well as the importance of an altered tau isoform ratio

caused by FUS variants. For this purpose, a previously established kinesin-1-dependent

microtubule gliding assay was modified to run reliably in the presence whole cell lysates, so that

the kinesin-1 motility can be monitored in a minimal system. In this assay, stabilized and

fluorescently labelled microtubules are propelled by a layer of underlying, surface bound

kinesin-1 motor proteins (Figure 7A). The collective motility of kinesins, however, depends on a

variety of external factors such as temperature, ionic strength, the presence of microtubule or

motor interacting proteins, or whole cell lysates. To study the impact of FUS variants or different

ratios of tau on this machinery, recombinant FUS-GFP and tau-GFP protein or whole cell

lysates of isogenic iPSC-derived spinal motor neurons with either wildtype FUS-GFP or

FUS-P525L-GFP are added to the assay (Figure 7B).

Figure 7: Reconstituting axonal transport in vitro.



A) ALS-patient-derived and isogenic control iPSC lines were differentiated towards spinal motor neurons. On day 21 of maturation, cells were mechanically lysed and centrifuged to obtain the soluble fraction of cell lysates, further referred to as whole cell lysates. B) Depending on the experiment, whole cell lysates or recombinant human protein (FUS-GFP or tau-GFP) were applied to a kinesin-1-dependent microtubule gliding assay in manually assembled flow chambers (see materials and methods for details).

To achieve efficient cell lysis and a high protein yield in the lysates, different lysis methods

have been tested (Table 9). Chemical lysis using the Active Motif Nuclear Extract Kit and a

variety of mechanical lysis protocols were assessed for their efficiency in terms of protein yield.

PBS was used in herein assessed mechanical lysis protocols as a preliminary lysis buffer, since

microtubule gliding velocity was as efficient in the presence of this physiological buffer as in the

presence of the most commonly used BRB80 buffer (Figure 8B) (Nitzsche et al., 2010). In

addition, two methods for determining the total protein concentration were evaluated for their

accuracy using lysates produced by chemical lysis, one based on the detection of

BCA/copper-complexes and one modified Bradford assay based on the direct binding of

Coomassie blue to proteins (Table 9B). In general, higher quantities of total protein were

detected using the BCA detection kit than with the Bradford detection. Importantly, the average

relative error for measurements performed with the Bradford assay was much larger

(0.078 ± 0.035) than for those performed with the BCA assay (0.013 ± 0.005), and hence total

protein levels in all mechanically prepared lysates were only quantified using the latter.

Previously established and widely used methods for mechanical lysis, such as glass bead

shearing, dounce homogenization, and liquid nitrogen grinding (Mackall et al., 1979; Harlow and

Lane, 2006; Walker, 2009), resulted in low protein yields as determined by BCA or did not

results in sufficient cell lysis, since the detected amount of the housekeeping protein β-actin was

very low as determined by western blot (Table 9A). A high yield, together with sufficient

detection of the housekeeping protein and hence cell lysis, was accomplished by shearing the

cells via passage through a syringe needle with a diameter of 400 µm or by dounce

homogenization. Although dounce homogenization yielded similar or even higher protein

concentrations, a lot of lysate stuck to the walls of the glass tube, so that overall lower volumes

of lysate were collected. This was not the case for mechanical lysis using a syringe and needle,

since all lysate could be dispensed with the plunger.



Table 9: Comparison of different protein isolation techniques for their cell lysis efficiency, determined by the detection of β-actin via western blot, and protein yield, determined by BCA or Bradford assay.

Representative images and values of single measurements. rel. err. = average relative error of the six depicted representative measurements ± standard deviation.

The highest protein yields in combination with sufficient protein lysis were obtained using

the chemical lysis kit (Table 9). However, microtubules frequently detached in the presence of

chemical lysis buffer, even after 10-fold dilution (Figure 8A). To circumvent this problem, the

buffer was exchanged by passing lysates through a PD MinitrapTM desalting and buffer

exchange column and eluting protein in PBS through gravity flow. This procedure resulted in a

severe loss of protein (Table 9B) due to the combination of protein being stuck to the column

surface and a high dilution of the lysate. In addition, a control experiment, where only chemical

lysis buffer was passed through the column and one elution step performed with PBS, did not

eliminate microtubule detachment entirely (Figure 8A), indicating that buffer exchange was not

sufficient to remove all substances interfering with microtubule-motor interaction. Similarly, while

buffer exchange lysate concentration using Vivaspin® sample concentrators resulted in very

high protein yield (Table 9A), the buffer could not be sufficiently exchanged using these columns

to prevent microtubule detachment (Figure 8A).



Therefore, whole cell lysates used in the herein presented study were prepared by

shearing cells in PBS via passage through a syringe needle, as this presented the most efficient

mechanical lysis method. Lysates were supplemented with a commercially available protease

inhibitor cocktail to prevent protein degradation. Using a commercially available phosphatase

inhibitor cocktail, however, immediately led to diminished microtubule gliding (Figure 8C).

Therefore, β-glycerophosphate was added to the lysate to steer phosphatase activity towards

the excess substrate. In addition, 10 % glycerol was added as a cryoprotectant. Glycerol by

itself did not alter microtubule gliding characteristics, but the addition of

10 mM β-glycerophosphate caused microtubules to detach from surface-bound motors (Figure

8D). To prevent this, 0.3 % methylcellulose was added to the assay to allow cross-comparison

as well to experiments involving recombinant proteins. The addition of methylcellulose did not

affect microtubule gliding, but effectively eliminated microtubule detachment (Figure 8D).

Methylcellulose due to its high viscosity acts as a cushion to keep microtubules close to the

imaging surface. Since microtubules sporadically detach from the surface-bound motors,

methylcellulose facilitates fast microtubule reattachment to motors (Inoue et al., 2015; Saito et

al., 2017). Cells were hence lysed in the presence of PBS and lysates supplemented with a

commercially available protease inhibitor, 10 % glycerol, 10 mM β-glycerophosphate, and

0.3 % methylcellulose (further referred to as lysis buffer). Similarly, standard motility buffer used

in subsequent microtubule gliding assays was prepared by addition of 10 µM taxol, 1 mM ATP,

20 mM D-glucose, 20 μg/mL glucose oxidase, 10 μg/mL catalase, 10 mM DTT, and

0.2 mg/mL casein to the lysis buffer.



Figure 8: Optimal buffer conditions for reproducible kinesin-1-dependent microtubule gliding in the presence of whole cell lysates consist of 10 % glycerol, 10 nM β-glycerophosphate and 0.3 % methylcellulose in PBS. Top of each panel: rhodamine-labelled microtubules gliding on surface-immobilized kinesin-1 in the presence of

(A) whole cell lysis buffer obtained from a Nuclear Extraction Kit for whole cell lysates (Active Motif) undiluted or at a 10-fold dilution, (B) PBS or BRB80 alone, (C) PBS supplemented with a commercially available phosphatase inhibitor, (D) PBS supplemented with 10 % glycerol, an additional 10 mM β-glycerophosphate, or moreover with 0.3 % methylcellulose. Yellow arrows indicate detaching microtubules. Scale bar = 10 µm. Bottom of each panel:

kymographs showing representative kinesin-1-dependt microtubule movement in the respective conditions. Horizontal scale bar = 2 µm, vertical = 5 s. E) Kinesin-1-dependent microtubule gliding velocities of six technical replicates in standard motility buffer. Relative velocities were calculated by dividing the microtubule gliding velocity after flushing with standard motility buffer by the microtubule velocity in the same channel before flushing.



Having optimized the buffer conditions, microtubule gliding assays were set up using

manually assembled flow chambers with six individual flow channels separated and tightly

sealed by stripes of parafilm in between two differently sized glass coverslips (Figure 6). Flow

channels were first perfused with a casein-containing solution to prevent unspecific protein

adsorption. This was followed by perfusion with a solution containing full-length constitutively-

active Drosophila melanogaster kinesin-1 motors. Both solutions were allowed to settle within

the channels at RT for 5 min each, before rhodamine-labelled microtubules in standard motility

buffer were flushed through the channels. Because the microtubule gliding velocity is highly

sensitive to temperature changes (Böhm et al., 2000), flow chambers were mounted onto a

custom-built Peltier element to minimize temperature fluctuations during an experiment and

between single technical replicates. Flow chambers were imaged using

widefield-epifluorescence microscopy with 10 s time lapse movies (Figure 9A, B) with an

exposure time of 100 ms and at an acquisition rate of one frame per second. From these time

lapses, frame-to-frame velocities of gliding microtubules were calculated using a previously

developed automated filament tracking algorithm (FIESTA, Ruhnow et al., 2011). The median

microtubule gliding velocity, as well as the 25th and 75th percentile (Figure 9C), were calculated

for microtubules in a given condition (i.e. presence of standard motility buffer, recombinant

protein, or whole cell lysates). Normalizing the median velocity to the initial median velocity (i.e.

before addition of recombinant FUS-GFP variants, recombinant tau isoforms, BSA, or standard

motility buffer) of microtubule in the same channel allows for cross-comparison of different

conditions and takes account of small experiment-to-experiment differences in e.g. the motor

density or motor activity. This normalization procedure further reduces the error arising from

possible fluctuations in microtubule number between experiments or technical replicates. With

this multitude of optimization steps, a robust assay to model axonal transport in vitro could be

established that generated highly reproducible measurements (Figure 9D).



Figure 9: Imaging and analysis of kinesin-1-dependent microtubule gliding velocity . A) Top panels: microtubule imaging was performed using widefield-epifluorescence microscopy to acquire time lapses of 10 s with an exposure time of 100 ms and an acquisition rate of one frame per second. Scale bar = 5 µm. Bottom panel: a kymograph representing the kinesin-1-dependent movement of the above microtubule. Horizontal

scale bar = 2 µm, vertical = 5 s. B) An overlay image illustrating the kinesin-1-dependent microtubule gliding in the presence of cell lysate containing wildtype FUS-GFP. Positions of microtubules at time points t = 0 s (cyan) and t = 10 s (magenta) are overlaid. White color in the overlay indicates positional overlap at both time points, and hence



no microtubule movement. Scale bar = 10 µm. C) A previously developed automated MATLAB script was used to calculate frame-to-frame velocities from 10 s time lapse movies. Frame-to-frame velocities of all time lapse movies in a given condition (e.g. cell lysate containing wildtype FUS-GFP) of one technical replicate were combined in a single histogram. The red solid line marks the median, while dashed red lines mark the 25

th and 75

th percentile, respectively.

D) Flow channels containing gliding rhodamine-labelled microtubules in standard motility buffer were flushed with buffer to rule out that the process of flushing solutions through a channel changes microtubule gliding velocity. Relative speed of gliding microtubules was calculated by dividing the microtubule gliding velocity after flushing with standard motility buffer by the velocity of microtubules in the same flow channel before flushing. Six technical replicates are shown.

3.2 Determination of assay sensitivity with the microtubule-associated

protein tau

In order to estimate the sensitivity of this modified kinesin-1-dependent microtubule gliding

motility assay for proteins interfering with motor motility on microtubules, human recombinant

full length (2N4R) tau-GFP was added to the assay at varying concentrations (Figure 10A). Tau

is a MAP known to inhibit kinesin-1 stepping on microtubules by a simple road-block mechanism

(Tarhan et al., 2013), thereby significantly reducing kinesin-1 run-length and velocity (Hoeprich

et al., 2014). To see whether tau shows the same behavior in our modified microtubule gliding

assay, recombinant 2N4R tau-GFP was added at final concentrations of 2 to 500 nM in

standard motility buffer, and the gliding velocity as well as microtubule binding were assessed

by widefield-epifluorescence microscopy. No significant binding of 2N4R tau-GFP to

microtubules was evident at low tau concentrations, while binding was observed at a

concentration of 50 nM and above (Figure 10B). The strong background signal of unbound tau-

GFP, however, prevented quantification of the exact amount of microtubule-bound tau, and

therefore binding was assessed only qualitatively.

In line with the observed microtubule-binding behavior of tau, kinesin-1-dependent

microtubule gliding velocity was not altered below a tau-GFP concentration of 24 nM, but

microtubules considerably slowed down in the presence of 59 nM 2N4R tau-GFP and

completely ceased movement at concentrations above 118 nM (Figure 10C). Hence, the

modified kinesin-1-dependent microtubule gliding assay is sensitive enough to detect the

presence of 60 nM 2N4R tau-GFP, which is equivalent to 2.65 µg/ml.



Figure 10: Tau-GFP binds to microtubules and inhibits kinesin-1-dependent microtubule gliding in a concentration dependent manner.

A) Full-length 2N4R recombinant human tau-GFP was introduced to a kinesin-1-dependent microtubule gliding assay in standard motility buffer. B) Binding of 2N4R tau-GFP (right panels) at indicated concentrations to rhodamine-labelled microtubules (left panels). Scale bar = 5 µm. C) Kinesin-1-dependent microtubule gliding velocities in the presence of 2N4R tau-GFP at indicated concentrations. Relative speed was calculated by dividing the median microtubule gliding velocities after tau-GFP addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before tau-GFP addition. Averages of three independent experiments ± standard deviation are shown.

3.3 Investigation of a direct interference of recombinant FUS-GFP with

kinesin-1 motility and microtubule gliding

Having established and verified the sensitivity of the herein modified kinesin-1-dependent

microtubule gliding assay for motor-microtubule interfering proteins, it was utilized to address

the fundamental question whether FUS variants directly interfere with the kinesin-1-mediated

transport machinery by e.g. binding to microtubules or inhibiting kinesin-1 motility (Figure 11A).

For this purpose, three increasing concentrations (5 nM, 500 nM, and 5000 nM) of recombinant

human wildtype FUS-GFP or FUS-P525L-GFP were added to the modified microtubule gliding

assay, and microtubule gliding as well as FUS-GFP binding to microtubules was again observed

using widefield-epifluorescence microscopy. Of note, the buffer control in these experiments

consisted of standard motility buffer supplemented with FUS-GFP protein storage buffer in

dilutions equal to those required to obtain the tested FUS-GFP concentrations, in order to

evaluate whether the storage buffer itself influences kinesin-1-dependent microtubule gliding.

Further, to observe whether kinesin-1 motility on microtubules is impaired in a crowded



environment due to the presence of high protein concentrations, as has previously been

reported (Sozański et al., 2015; VanDelinder et al., 2020), equimolar amounts of BSA were

tested in the kinesin-1-dependent microtubule gliding assay as an additional control.

Neither FUS-GFP variant showed any specific microtubule binding (Figure 11B). In line

with this, kinesin-1-dependent microtubule gliding was also not significantly affected by the

addition of either FUS-GFP variant in comparison to experiments with equal concentrations of

BSA as a protein crowding control or to a buffer control (Figure 11C, Table S 1). Since no

inhibitory effects were found, it can be concluded that neither wildtype FUS-GFP nor

FUS-P525L-GFP directly binds to microtubules nor inhibits kinesin-1 transport motility over a

wide range of concentrations.

Figure 11: Recombinant FUS-GFP variants to not inhibit kinesin-1-dependent microtubule gliding.

A) Recombinant human FUS-GFP variants were introduced to a kinesin-1-dependent microtubule gliding assay in standard motility buffer. B) Binding of wildtype FUS-GFP or FUS-P525L-GFP (right panels) at indicated concentrations to rhodamine-labelled microtubules (left panels). Scale bar = 5 µm. C) Kinesin-1-dependent microtubule gliding velocities in the presence of wildtype FUS-GFP, FUS-P525L-GFP, or BSA in standard motility buffer at indicated concentrations. Relative speed was calculated by dividing the median microtubule gliding velocities after FUS-GFP or BSA addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before FUS or BSA addition, respectively. Averages of three independent experiments ± standard deviation are shown.



This study focused on the interference of free-in-solution, non-aggregated FUS variants

with kinesin-1 motility on microtubules. In vivo, cytoplasmic mislocalized mutant FUS variants

form aggregates (Patel et al., 2015) over the course of hours, developing from liquid

compartments in the cytoplasm that undergo a liquid-to-solid phase transition (Mateju et al.,

2017; Nötzel et al., 2018). These aggregation events can be reproduced to some extent in vitro,

where they occur at a time scale of seconds in buffer with high ionic strength (Marrone et al.,

2019). Buffers used in the present study were of low to moderate ionic strength, and hence

FUS-GFP aggregate formation was, as expected, not observed directly after the addition of

FUS-GFP variants to the kinesin-1-dependent microtubule gliding assay (Figure 11B). However,

it cannot be excluded that – similar to the in vivo situation – FUS-GFP variants aggregate over

time and may further only in this state exhibit direct inhibitory effects on kinesin-1 motility.

Therefore, kinesin-1-dependent microtubule gliding assays were repeated as above, but the

microtubule binding and gliding velocity in the presence of wildtype FUS-GFP or

FUS-P525L-GFP was now observed over the course of three hours. Although the microtubule

gliding velocities in the presence of either FUS-GFP variant declined linearly over time, this

effect can be attributed to assay ageing since all control experiments (i.e. presence of standard

motility buffer or BSA at equivalent concentrations) showed the same age-dependent decline in

microtubule gliding velocity (Figure 12A, Table S 1). This is again in line with the finding that

neither FUS-GFP variant bound to microtubules over the course of three hours, and neither

could aggregate formation be observed in the presence of those FUS-GFP variants during the

same time course (Figure 12B). From both experiments can hence be concluded that neither

wildtype FUS-GFP nor the FUS-P525L-GFP variant in its non-aggregated form interferes with

kinesin-1 motility on or binds to microtubules, even over the course of hours.



Figure 12: FUS-GFP variants do not inhibit kinesin-1-dependent microtubule gliding velocity and do not aggregate over the course of three hours.

A) Kinesin-1-dependent microtubule gliding velocities over the course of three hours in the presence of indicated concentrations of wildtype FUS-GFP, FUS-P525L-GFP, protein control (BSA), or standard motility buffer. Relative speed was calculated by dividing the median microtubule gliding velocities after FUS-GFP addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before FUS-GFP addition. Averages of three independent experiments ± standard deviation are shown. B) Recombinant human wildtype FUS-GFP and FUS-P525L-GFP in standard motility buffer at indicated concentrations and time points. Scale bar = 10 µm.



3.4 Kinesin-1-dependent microtubule gliding and assay sensitivity in the

presence of whole cell lysates expressing GFP-labelled FUS variants

FUS-GFP variants did not directly interfere with kinesin-1-dependent microtubule gliding (e.g. by

binding to microtubules or motors). However, kinesin-1 often requires a multitude of adaptor

proteins in order to efficiently bind its cargo (Hirokawa et al., 2009; Fu and Holzbaur, 2014).

Using whole cell lysates from spinal motor neurons expressing GFP-labelled FUS variants in the

modified kinesin-1-dependent microtubule gliding assay, we aimed to investigate whether FUS

requires endogenous adaptors or whether there are indirect effects (e.g. changes in expression

levels of other motor or microtubule effector proteins (Fujioka et al., 2013; Akiyama et al., 2019)

or sequestering of MAPs etc. in aggregates (Yasuda et al., 2017) that may impact the

kinesin-1-dependent transport on microtubules. Therefore, previously generated and

characterized ALS-patient-derived and CRISPR/Cas9-engineered isogenic iPSCs (Naumann et

al., 2018; Pal et al., 2018) were differentiated via NPCs to form spinal motor neurons expressing

either wildtype FUS-GFP or FUS-P525L-GFP. Whole cell lysates were obtained after 21 days of

neuron maturation by mechanical lysis through a thin needle. Lysates were cleared by

centrifugation in order to obtain only soluble parts within the lysate, especially free floating,

cytoplasmic FUS. For application of whole cell lysates in the kinesin-1-dependent microtubule

gliding assay (Figure 13A), the total protein concentration in the lysates was determined using

the BCA assay, and lysates were diluted in cell lysis buffer to a final concentration of

50, 80, 110, and 140 ng/µL. Due to low yields after protein isolation, higher concentrations could

not be tested. Imaging and analysis was performed as described above, with respective

concentrations of BSA in standard motility buffer again serving as a protein crowding control.

However, neither lysates from wildtype FUS-GFP nor FUS-P5525L-GFP expressing cells had

any significant effect on microtubule gliding compared to controls (Figure 13B, Table S 3).



Figure 13: Cell lysates of spinal motor neurons expressing ALS-associated or wildtype FUS variants do not interfere with microtubule gliding on kinesin-1 motors.

A) Lysates of cells expressing wildtype FUS-GFP or FUS-P525L-GFP were applied to the modified kinesin-1-dependent microtubule gliding assay. B) Kinesin-1-dependent microtubule gliding velocities in the presence of cell lysates in which wildtype FUS-GFP and FUS-P525L-GFP was expressed, respectively, or BSA in standard motility buffer at indicated concentrations. Relative speed was calculated by dividing the median microtubule gliding velocities after lysate or BSA addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before lysate or BSA addition, respectively. Averages of four independent experiments ± standard deviation are shown, with n = 4. C) Kinesin-1-dependent microtubule gliding velocities in the presence of cell lysates in which wildtype FUS-GFP and FUS-P525L-GFP was expressed, respectively, or BSA in standard motility buffer at a concentration of 80 ng/µL supplemented with indicated concentrations of recombinant human 2N4R tau-GFP. In addition, titration of 2N4R tau-GFP from Figure 10C is included for comparison. Relative speed was calculated by dividing the median microtubule gliding velocities after lysate, BSA, or tau-GFP addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before lysate, BSA, or tau-GFP addition, respectively. Averages of three independent experiments ± standard deviation are shown.

In order to estimate the amount of FUS-GFP variants present within cell lysates and

compare it to the concentrations of recombinant FUS-GFP variants tested above, a western blot

was performed with four biological replicates of cell lysates expressing wildtype FUS-GFP or

FUS-P525L-GFP, respectively (Figure 14). The level of endogenous wildtype FUS-GFP and

FUS-P525L-GFP protein within the herein used cell lysates was much lower (between 2.5 and

5 nM) than the amount of recombinant human FUS-GFP variants used in the above described

experiments. Therefore, it is unlikely that potential direct interactions of FUS-GFP variants in cell

lysates with kinesin-1 or microtubules were missed due to the low concentration of endogenous



protein, as recombinant FUS-GFP variants were supplied at a wide range of much higher

concentrations (5-5000 nM) and did not interfere with kinesin-1-dependent microtubule gliding.

Figure 14: Western blot of FUS-GFP variants expressed in whole cell lysates of spinal motor neurons.

Representative western blot of recombinant wildtype FUS-GFP at indicated concentrations and four biological replicates (n) of whole cell lysates obtained from spinal motor neurons expressing either wildtype FUS-GFP or FUS-P525L-GFP.

Due to the protein-rich environment in cell lysates, however, the sensitivity of the

kinesin-1-dependent microtubule gliding assay might be reduced. Hence, to test assay

sensitivity, recombinant human 2N4R tau-GFP was titrated at low amounts (2 – 237 nM) into the

lysates (which were diluted to a total protein concentration of 80 ng/µL). Assay performance was

re-evaluated to see if a slowdown of microtubule gliding velocity could still be detected below a

concentration of 118 nM tau-GFP in the presence of whole cell lysates. In contrast to

experiments with tau-GFP alone, kinesin-1-dependent relative microtubule gliding velocity

started to decrease already in the presence of 12 nM 2N4R tau-GFP in lysates, but also

completely ceased at a concentration of 118 nM substituted tau-GFP (Figure 13C), similar to

what was observed in the absence of cell lysates. This behavior was also independent of the

FUS variant expressed within the lysate. This implies that, even in a protein-rich environment

such as in the presence of whole cell lysates, the assay is still sufficiently sensitive to inhibitors

of kinesin-1 motility. Further, it can be concluded that neither lysates carrying wildtype FUS-GFP

nor those containing ALS-associated FUS-P525L-GFP negatively affect kinesin-1 motility.

3.5 Arrest in gliding of single microtubules caused by Tau-GFP compared

to aging of the assay

Since aging of the assay over the course of three hours and the presence of 2N4R tau-GFP

both reduced the relative microtubule gliding velocity to a similar extent (Figure 10 and Figure

12), the mode of action by which this slowdown occurred was further investigated. In the



presence of standard motility buffer, microtubules of one technical replicate glide with

frame-to-frame velocities around 600 nm/s, while microtubule gliding was entirely arrested in the

presence of 118 nM 2N4R tau-GFP (Figure 15A). When 24 nM 2N4R tau-GFP are

supplemented to the assay, microtubules display two heterogeneously moving populations, one

non-gliding fraction indicated by a peak in frame-to-frame velocities around zero, and one

fraction of microtubules that are gliding with frame-to-frame velocities broadly distributed

between 200 and 800 nm/s, resulting in a median frame-to-frame velocity of about 250 nm/s

(Figure 15A). A similar median frame-to-frame velocity of about 300 nm/s could be observed in

the presence of 5000 nM recombinant FUS-P525L-GFP two hours after protein administration,

although the peak of frame-to-frame velocities around zero was much smaller. Rather, most

microtubule frame-to-frame velocities lay between 100 and less than 600 nm/s (Figure 15A).

This indicates that microtubules collectively glide at a reduced frame-to-frame velocity when the

assay is aging, possibly due to subsiding motility of the underlying kinesin-1 motors, while the

presence of small amounts of tau-GFP causes an arrest in gliding of individual microtubules

while others are still propelled at about similar speed as in standard motility buffer. In order to

investigate this hypothesis further, the fraction of non-gliding microtubules in the presence of

2N4R tau-GFP, FUS-GFP variants, cell lysate or BSA, respectively, was calculated as the

number of microtubules gliding at a velocity less than 20 % of the median microtubule velocity in

standard motility buffer, and normalized to the total number of microtubules in one time-lapse

movie (Figure 15B-F). In the presence of standard motility buffer, the fraction of non-gliding

microtubules consistently appears around 0.1. Corresponding to a decrease in relative

microtubule gliding velocity upon 2N4R tau-GFP addition (Figure 10C), the fraction of

non-gliding microtubules begins to increase in the presence of 24 nM 2N4R tau-GFP and

continues to grow with increasing tau-GFP concentration until virtually all microtubules have

stopped gliding above 118 nM tau-GFP (Figure 15B). In contrast, the fraction of non-gliding

microtubules remained around 0.1 in the presence of either FUS-GFP variant at a concentration

of 5, 500, or 5000 nM, or equal concentrations of BSA (Figure 15C). Even over the course of

three hours, the fraction of non-gliding microtubules under these conditions only slightly

increased to a maximum of 0.20 ± 0.04 (i.e. for 5000 nM wildtype FUS-GFP after three hours,

Figure 15D). Similarly, the presence of cell lysates where either wildtype FUS-GFP or

FUS-P525L-GFP was expressed resulted in a low fraction of non-gliding microtubules around

0.1, independent of the total protein concentration (Figure 15E). When varying concentrations of

2N4R tau-GFP were added to either cell lysates, however, the fraction of non-gliding

microtubules started to increase already at a concentration of 12 nM supplemented tau-GFP,



and again continuously rose until virtually all microtubules have stopped gliding at a

concentration of 118 nM 2N4R tau-GFP or above (Figure 15F).

Figure 15: 2N4R Tau-GFP binding to microtubules causes an arrest of microtubule gliding.

A) Representative frame-to-frame velocities of one technical replicate of microtubules gliding in the presence of standard motility buffer, 2N4R tau-GFP, or FUS-P525L-GFP at the indicated concentrations and time points. Red



solid lines mark the median, while dashed red lines mark the 25th and 75

th percentile, respectively. B-F) The fraction

of non-gliding microtubules describes the number of microtubules in one technical replicate gliding with a frame-to-frame velocity less than 20 % of the gliding velocity in standard motility buffer, normalized to the total number of microtubules in one time lapse movie, before the addition of B) 2N4R tau-GFP at the indicated concentrations, C) recombinant human wildtype FUS-GFP, FUS-P525L-GFP, or BSA, at the indicated concentrations, D) standard motility buffer, recombinant human wildtype FUS-GFP, FUS-P525L-GFP, or BSA at the indicated concentrations over the course of three hours, E) cell lysates expressing wildtype FUS-GFP or the FUS-P525L-GFP variant, or BSA, at the indicated concentrations, and F) cell lysates expressing wildtype FUS-GFP or the FUS-P525L-GFP variant, or BSA at a concentration of 80 ng/µL supplemented with 2N4R tau-GFP at the indicated concentrations.

Of note, while the relative microtubule gliding velocity in the presence of recombinant

wildtype FUS-GFP decreases over the course of three hours down to 0.65 ± 0.14 (Figure 12A),

the fraction of non-gliding microtubules in this condition only increases up to 0.20 ± 0.04 within

the same time frame (Figure 15D). In contrast, the relative microtubule gliding velocity in the

presence of 35 nM and 59 nM 2N4R tau-GFP decreases from 0.80 ± 0.27 to 0.25 ± 0.35,

respectively (Figure 10C), accompanied by a drastic increase in the fraction of non-gliding

microtubules from 0.26 ± 0.14 to 0.72 ± 0.10, accordingly (Figure 15B). Interestingly, the relative

microtubule gliding velocity tends to decrease stronger when cell lysates supplemented with

2N4R tau-GFP are administered to gliding microtubules compared to equal concentrations of

tau-GFP alone (Figure 13C), while the fraction of non-gliding microtubules correspondingly also

tends to increase at lower concentrations of tau-GFP supplemented to cell lysates (Figure 15F).

Together, these findings suggest that 2N4R tau-GFP causes an arrest in the gliding of individual

microtubules, while others seem to be propelled with almost equal velocities as in standard

motility buffer. Although tau-decoration on microtubules often corresponded to arrest of

microtubule gliding, some tau-decorated microtubules were still propelled by underlying

kinesin-1 motors in the presence of low amounts of tau-GFP, e.g. 35 nM (Figure 16), indicating

that microtubule gliding is only reduced when a threshold amount of tau binds to microtubules

and further declines the more tau attaches to the filament.

Figure 16: 2N4R Tau-GFP binding to microtubules at low concentrations leads to an arrest in gliding of individual microtubules, while others are still motile.

A) Binding of2N4R tau-GFP at a concentration of 35 nM to rhodamine-labelled microtubule. Scale bar = 5 µm. B) Top panels: rhodamine-labelled microtubules gliding (1) or non-gliding (2) in the presence of 35 nM 2N4R tau-GFP. Scale bar = 5 µm. Bottom panels: kymographs representing the kinesin-1-dependent movement of the above gliding (1) or non-gliding (2) microtubule. Horizontal scale bar = 2 µm, vertical = 5 s.



3.6 Differential effects of 2N3R and 2N4R tau individually on

kinesin-1-dependent microtubule gliding velocity and microtubule binding

While this work shows that FUS variants have no direct effect on kinesin-1-dependent transport,

they might indirectly disturb axonal transport by changing the equilibrium of tau isoforms in the

axon due to nuclear loss-of-function of FUS-P525L (Fujioka et al., 2013; Ishigaki et al., 2017).

Under physiological conditions, FUS directly binds to tau pre-mRNA in the nucleus and,

together with the splicing factor SFPQ, regulates its alternative splicing by skipping exon 10,

resulting in 3R tau isoforms (Orozco et al., 2012; Ishigaki et al., 2017; Ishigaki and Sobue,

2018). This is in line with the observation that upon FUS mislocalization to the cytoplasm, exon

10 is more frequently incorporated into the mature tau-mRNA, which results in an increased

translation of 4R tau isoforms. The increase in 4R:3R isoform ratio has previously been

indicated to cause a variety of pathological changes within the cell, such as slightly shorter

axons and highly enlarged growth cones with less bundled and further spread microtubules

(Orozco et al., 2012). This toxic effect on neurite outgrowth could be rescued by silencing 4R

tau (Ishigaki et al., 2017). This indicates how detrimental a shift in 4R:3R tau isoform ratio can

be for the physiological function of the cell. However, it is not known to date if and to what

extend the shift in 4R:3R tau isoform ratio, caused by the cytoplasmic mislocalization of FUS-

P525L, contributes to the axonal transport defects observed in ALS pathology.

This study therefore aimed at investigating the direct effect of 2N4R and 2N3R tau

isoforms, alone or in combination, on the kinesin-1-dependent microtubule gliding. For this

purpose, recombinant human 2N3R tau-GFP and 2N4R tau-mScarlet labelled isoforms were

expressed in and purified from insect cells (see 2.2.7). Each tau variant was at first added

individually to the modified kinesin-1-dependent microtubule gliding assay at increasing

concentrations (Figure 17A and B), and microtubule gliding again observed using widefield-

epifluorescence microscopy. BSA was again used as a protein crowding control at the highest

concentration of tau variants tested (5.25 µM). While the relative microtubule gliding velocity

goes down with increasing concentrations of either isoform (Figure 17C) compared to the

microtubule gliding in standard motility buffer or BSA, the 4R tau-mScarlet isoform causes a

slowdown of microtubule gliding already at much lower concentrations (starting at 0.05 µM tau)

compared to the 3R tau-GFP isoform. At 0.8 µM 4R tau-mScarlet, the relative microtubule

gliding speed has decreased to 0.31 ± 0.08, while the 3R tau-GFP isoforms causes a similar

drop of relative microtubule gliding speed only at a more than 5-fold higher concentration

(5.25 µM).



Figure 17: The 2N4R tau isoform interferes with microtubule gliding on kinesin-1 motors at much lower concentrations compared to the 2N3R tau isoform.

A) Recombinant human 2N3R tau-GFP was introduced to a kinesin-1-dependent microtubule gliding assay in standard motility buffer. B) Recombinant human 2N4R tau-mScarlet was introduced to a kinesin-1-dependent microtubule gliding assay in standard motility buffer. C) Kinesin-1-dependent microtubule gliding velocities in the presence of recombinant human 2N3R tau-GFP, 2N4R tau-mScarlet, at indicated concentrations, or BSA in standard motility buffer at a concentration of 5.25 µM. Relative speed was calculated by dividing the median microtubule gliding velocities after tau variant or BSA addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before tau variant or BSA addition, respectively. Averages of three independent experiments ± standard deviation are shown.

Since the two tested isoforms differ mainly in their number of microtubule binding

domains, their different capacity to slowdown kinesin-1-dependent microtubule gliding might be

explained by their distinct microtubule binding affinities. To test this hypothesis, each tau variant

was added individually to taxol-stabilized, surface-immobilized, and Atto647-labelled

microtubules at increasing concentrations, and the amount of tau bound to microtubules was

observed using TIRF microscopy (Figure 18A and B).

Binding of the 2N3R tau-GFP isoform to microtubules was apparent at 100 nM tau in

solution, while saturation was achieved at 800 nM tau in solution (Figure 18A). In line with the

observed slowdown of gliding microtubules in the presence of low concentrations of

2N4R tau-mScarlet, this tau isoform bound to microtubules already at low concentrations

(i.e. 5 nM, Figure 18B), while saturation was already achieved at 200 nM.



The fraction of tau protein that was bound to microtubules when a specific concentration

of tau was present in solution, relative to the amount of the respective tau isoform bound at the

highest concentration tested, was plotted and the data fitted using the Hill equation (Hill, 1910)

in order to describe the binding kinetics of the respective tau isoform (Gesztelyi et al., 2012)

(Figure 18C and D). The respective value for the highest concentration tested for each tau

isoform (800 nM for 3R tau-GFP and 200 nM for 4R tau-mScarlet, respectively) was excluded

from the fit due to the fact that it was used for normalization and yielded a standard error of the

mean (SEM) of zero after averaging the experiments. Interestingly, the Hill equation determined

a Hill coefficient (n) larger than 1 for both binding curves for 3R tau-GFP and 4R tau-mScarlet,

namely 2.38 ± 0.67 and 1.83 ± 0.29, respectively. This indicates a cooperative binding

mechanism for both tau isoforms (Weiss, 1997). The dissociation constant Kd (or k in Figure

18C and D), corresponding to the concentration at which half of the tau molecules in solution

are bound to microtubules, for 3R tau-GFP (Kd = 137.12 ± 29.7 nM) was determined to be

approximately 20-fold higher compared to 4R tau-mScarlet (Kd = 6.79 ± 0.41 nM), indicating that

the binding affinity of 4R tau-mScarlet to microtubules is approximately 20-fold higher than that

of 3R tau-GFP.

Interestingly, although the microtubule was heavily decorated with 2N3R tau-GFP

molecules, the relative microtubule gliding speed at 750 nM 2N3R tau-GFP was only reduced to

0.85 ± 0.07 (Figure 17C). Similarly, microtubule saturation with 2N4R tau-mScarlet at 250 nM

corresponded to a decrease in relative kinesin-1-dependent microtubule gliding speed only

down to 0.48 ± 0.13 (Figure 17C). For both recombinant tau isoforms, the relative microtubule

gliding velocity decreased further at higher concentrations than those needed to saturate

microtubules with tau. We hence suggest that tau molecules do not just form a monolayer on

microtubules, but can cluster on the surface and impair kinesin-1 motility stronger than single

tau molecules.



Figure 18: The 2N4R tau-mScarlet isoform has a stronger binding affinity towards microtubules compared to 2N3R tau-GFP.

A) Binding of 2N3R tau-GFP (upper panels) at indicated concentrations to Atto647-labelled microtubules (lower panels). Scale bar = 5 µm. B) Binding of 2N34 tau-mScarlet (upper panels) at indicated concentrations to Atto647-labelled microtubules (lower panels). Scale bar = 5 µm. C) The fraction of bound tau describes the amount of tau bound to microtubules at the indicated concentration relative to the amount of tau bound at the highest tested concentration of 3R tau-GFP. At least 200 microtubules were analyzed for each concentration and in every experiment. Averages of three independent experiments ± SEM are shown. The last data point was excluded from Hill fit due to its SEM=0 after normalization. D) The fraction of bound tau describes the amount of tau bound to microtubules at the indicated concentration relative to the amount of tau bound at the highest tested concentration of 4R tau-mScarlet. At least 200 microtubules were analyzed for each concentration and in every experiment. Averages of three independent experiments ± SEM are shown. The last data point was excluded from Hill fit due to its SEM=0 after normalization.



3.7 Impact of increasing 4R:3R tau isoform ratios on kinesin-1-dependent

microtubule gliding and its microtubule binding

After having determined the effect of 2N3R tau-GFP and 2N4R tau-mScarlet on

kinesin-1-dependent microtubule gliding individually, we next tested whether a small shift in tau

isoform ratio towards 2N4R tau is sufficient to differentially affect kinesin-1-dependent

microtubule gliding. Previous studies reported that upon FUS knockdown, mimicking the

mislocalization of FUS-P525L from the nucleus to the cytoplasm, the ratio of total 4R:3R tau

isoforms within the cell changes from approximately 0.6 to 1.3 (Ishigaki et al., 2017). To

reconstitute this effect in vitro, both recombinant human 2N3R tau-GFP and 2N4R tau-mScarlet

isoforms were combined at increasing ratios with a constant total tau concentration of 1200 nM

and administered to the modified microtubule gliding assay (Figure 19A). Ratios in this study

were calculated according to the following formula and are summarized in Table 10:

[4𝑅 𝑡𝑎𝑢]

[3𝑅 𝑡𝑎𝑢 + 4𝑅 𝑡𝑎𝑢]= 1200 𝑛𝑀

Table 10: Calculation scheme for 4R:3R tau isoform ratios used in this study.

[𝟒𝑹 𝒕𝒂𝒖]

[𝟑𝑹 𝒕𝒂𝒖 + 𝟒𝑹 𝒕𝒂𝒖] = 𝟏𝟐𝟎𝟎 𝒏𝑴

Amount 4R

tau-GFP (nM)

Amount 3R

tau-mScarlet (nM) 4R:3R Ratio

0 0 1200 0

0.25 300 900 0.33

0.5 600 600 1

0.75 900 300 3

1 1200 0 (4)

Interestingly, the relative kinesin-1-dependent microtubule gliding speed decreases from

0.58 ± 0.1 in the presence of 4R tau-mScarlet and 3R tau-GFP at a ratio of 0.33, to a relative

microtubule gliding speed of 0.34 ± 0.19 in the presence of 4R tau-mScarlet and 3R tau-GFP at

a ratio of 3 (Figure 19B), compared to a buffer control or gliding in the presence of 3R tau-GFP

alone. This indicates that an increase in the ratio of 4R:3R tau isoforms, as subtle as observed

in vivo and without an increase in total tau levels, is sufficient to impair kinesin-1 motility on

microtubules. Hence, an increase in 4R:3R tau isoform ratio upon depletion of FUS-P525L from

the nucleus may contribute to the observed defects in axonal transport in FUS-ALS pathology.



Figure 19: An increase in 4R:3R tau isoform ratio leads to impaired kinesin-1-dependent microtubule gliding.

A) Recombinant human 2N3R tau-GFP was mixed at different ratios with 2N4R tau-mScarlet and introduced to a kinesin-1-dependent microtubule gliding assay in standard motility buffer. B) Kinesin-1-dependent microtubule gliding velocities in the presence of recombinant human 2N3R tau-GFP and 2N4R tau-mScarlet mixed at indicated ratios. Relative speed was calculated by dividing the median microtubule gliding velocities after tau mixture addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before tau mixture addition, respectively. Averages of three independent experiments ± standard deviation are shown.

In order to determine how increasing ratios of 4R:3R tau isoforms affect the microtubule

binding affinity of each isoform, both 4R tau-mScarlet and 3R tau-GFP were again mixed at the

same increasing ratios as for microtubule gliding experiments and added to taxol-stabilized,

surface-immobilized Atto647-labelled microtubules. The amount of tau bound to microtubules

was again observed using TIRF microscopy (Figure 20A) and quantified (Figure 20B).

Intriguingly, when 4R tau-mScarlet and 3R tau-GFP are supplied in combination to

surface-immobilized microtubules, 4R tau-mScarlet strongly bound to microtubules and inhibited

binding of 3R tau-GFP at a ratio as little as 0.33 (which is equal to 900 nM 3R tau-GFP and

300 nM 4R tau-mScarlet in solution). Further, the higher the 4R:3R ratio was in solution, the

less 3R tau-GFP was able to bind to microtubules until virtually none bound at a 4R:3R ratio of 3

(equal to 300 nM3R tau-GFP and 900 nM 4R tau-mScarlet in solution). Interestingly, at this

ratio, both tau isoforms were present at a concentration at which microtubules were saturated

when only one isoform was added to the assay individually (Figure 18). In addition, the

microtubule gliding velocity decreases at increasing 4R:3R tau ratios in a similar fashion as in

the presence of 2N4R tau-mScarlet alone. This indicates that i) 2N4R tau-mScarlet actively

replaces 2N3R tau-GFP on the microtubule and ii) 2N4R tau-mScarlet is more effective in

blocking kinesin-1 motility than 2N3R tau-GFP, since the total concentration of tau is equal at all

ratios.



Figure 20: The 2N4R tau-mScarlet isoform prevents binding of 2N3R tau-GFP to microtubules due to its stronger binding affinity.

A) Binding of 2N3R tau-GFP (middle panels) and 2N4R tau-mScarlet (lower panels) at indicated ratios to Atto647-labelled microtubules (uppermost panels). Scale bar = 5 µm. B) The fraction of bound tau describes the amount of tau bound to microtubules at the indicated concentration relative to the amount of tau bound at the highest tested concentration of the respective tau isoform. At least 200 microtubules were analyzed for each ratio and tau isoform in every experiment. Averages of three independent experiments ± standard deviation are shown.



3.8 Expression levels of 4R and 3R tau isoforms in whole cell lysates

The so far presented experimental data strongly points towards the fact that an increase in

4R:3R tau isoform ratio within the cell contributes to the observed defects in axonal transport.

This is likely due to a higher decoration of microtubules with 4R tau isoforms and their larger

roadblock effect compared to 3R tau isoforms. An increased 4R:3R ratio likely results from the

cytoplasmic mislocalization of the FUS-P525L mutant as observed in the used cell lines

(Dormann et al., 2010; Ishigaki et al., 2017). This would indicate that the 4R:3R tau isoform ratio

should be increased in whole cell lysates expressing the FUS-P525L-GFP variant compared to

lysates from cells expressing the wildtype FUS-GFP variant. However, when both lysates were

added to the herein used kinesin-1-dependent microtubule gliding assay, they did not

differentially influence the relative microtubule gliding speed (Figure 13). Therefore, western

blots were performed to determine the level of each tau isoform within the herein used whole

cell lysates (Figure 21). In order to detect 3R and 4R tau isoforms individually, primary

pan-specific antibodies detecting either 0N-, 1N-, and 2N3R tau isoforms or 0N-, 1N-, and 2N4R

tau isoforms were used in these blots. Recombinant human 2N3R tau-GFP and

2N4R tau-mScarlet was used as a positive control, respectively. Interestingly, none of the 4R

tau isoforms could be detected in four biological replicates of whole cell lysates expressing the

wildtype FUS-GFP or FUS-P525L-GFP variants, respectively (Figure 21B), as well as the 1N3R

and 2N3R tau isoforms (Figure 21A).The 0N3R tau isoform, however, was consistently present

in all biological replicates with a concentration between 10 and 30 nM. This indicates that the

expression level of tau isoforms in the herein used iPSC-derived spinal motor neurons has not

(yet) been influenced by the FUS-P525L-GFP variant, at least not within the detection limit of

this western blot technique. However, the presence of small amounts of 0N3R tau within lysates

might contribute to the non-significant, but consistently lower relative microtubule gliding speeds

observed in the kinesin-1-dependent microtubule gliding assay in the presence of whole cell

lysates supplemented with recombinant human 2N4R tau-GFP, compared to the presence of

2N4R tau-GFP alone (Figure 13C). Alternatively, unknown factors present in whole cell lysates

may interfere with kinesin-1-dependent microtubule gliding, which could be determined by e.g.

pulldown experiments with microtubules and motors in the presence of whole cell lysates and

subsequent mass spectrometry.



Figure 21: Western blot of tau variants expressed in whole cell lysates of spinal motor neurons.

Representative western blot of A) recombinant human 2N3R tau-GFP and b) recombinant human 2N4R tau-mScarlet

at indicated concentrations and four biological replicates (n) of whole cell lysates (at a concentration of 80 ng/µL)

obtained from spinal motor neurons expressing either wildtype FUS-GFP or FUS-P525L-GFP. The blot in A) was incubated with primary antibodies against 0N-, 1N-, and 2N3R tau isoforms, while the blot in B) was incubated with primary antibodies against 0N-, 1N-, and 2N4R tau isoforms.

Mechanisms of Axonal Transport in ALS Discussion


4. Discussion

The pathology of ALS has been subject to many previous studies, and yet there is no clear

consensus about the exact mechanisms leading to this devastating disease. Since mutations in

SOD1 were the first discovered in ALS patients, most research results to date are based on

models with this mutation (Rosen et al., 1993; Huai and Zhang, 2019), but intensive studies

have also been performed on the role of mutant FUS variants (Groen et al., 2010; Shang and

Huang, 2016; Naumann et al., 2019). Although defects in axonal transport have long been

observed in FUS-ALS (De Vos et al., 2008; Ikenaka et al., 2012; De Vos and Hafezparast,

2017b), little is known about underlying pathomechanisms that lead to these detrimental

defects, and whether they are causative or occur as a consequence of other pathological

events. The work done for this thesis therefore aimed at investigating the fundamental

hypothesis of a direct influence of NLS-mutant, cytoplasmic mislocalized FUS on the axonal

transport machinery involving kinesin-1. For this purpose, a kinesin-1-dependent microtubule

gliding assay was modified to be compatible with and robustly deliver reproducible results in the

presence of protein-rich solutions such as whole cell lysates. To evaluate the direct influence of

FUS variants on kinesin-1 motility, recombinant human GFP-labelled FUS variants, as well as

cell lysates from iPSC-derived spinal motor neurons expressing the same GFP-labelled FUS

variants, were analyzed in this assay. In addition, a more indirect effect of mutant FUS variants

on axonal transport was hypothesized and investigated: FUS is known to regulate splicing of

mRNA coding for the microtubule-binding protein tau, which suggests that the ratio of 4R:3R tau

is increased in cells carrying a mutation in FUS leading to its cytoplasmic mislocalization

(Ishigaki et al., 2017). Therefore, the modified kinesin-1-dependent microtubule gliding assay

was utilized to study whether and which 3R:4R tau ratio is sufficient to alter kinesin-1 motility on

microtubules.

This study therefore presents the first to address the direct role of FUS variants on

kinesin-1 motility in a bottom-up in vitro reconstruction of axonal transport. Likewise, using the

same reconstruction approach, it provides further evidence for the relevance of tau isoform

variations in disease progression. The results presented herein therefore largely contribute to

our understanding of the underlying pathomechanisms in FUS-ALS.



4.1 The modified kinesin-1-dependent microtubule gliding assay detects

nanomolar amounts of recombinant human tau-GFP

The buffer typically used in standard kineins-1-dependent microtubule gliding assays is BRB80

(with PIPES as a buffering agent), which we aimed to substitute with PBS for optimal cell lysis at

physiological pH (7.4). To prevent water from crystallizing and hence damage proteins during

snap freezing, glycerol was added to every cell lysate, as well as a commercially available

protease inhibitor cocktail. Since the phosphorylation state of proteins, in this case especially of

tau, is important for their activity, phosphatases within lysates needed to be inhibited as well.

However, a commercially available phosphatase inhibitor cocktail introduced to the microtubule

gliding assay led to an immediate arrest of microtubule gliding (Figure 8C). According to the

manufacturer, the tested phosphatase inhibitor cocktail was a mixture of the broad-spectrum

inhibitors sodium fluoride, sodium pyrophosphate, sodium orthovanadate and

β-glycerophosphate. Sodium fluoride and sodium pyrophosphate are general inhibitors for

phosphoseryl and phosphothreonyl phosphatases and commonly used in the purification of

protein kinases (Gordon, 1991; Hardie, 1993; Mercan and Bennett, 2010). The serine-threonine

phosphatase inhibitor β-glycerophosphate acts as a bait substrate (Belfield and Goldberg, 1968;

Chung et al., 1992; Lecanda et al., 1997), thereby protecting a protein’s phosphorylation status

by attracting phosphatases away from endogenous proteins. Sodium orthovanadate is a general

inhibitor for phosphotyrosyl phosphatases, alkaline phosphatases and a variety of ATPases

(Gordon, 1991; Wang et al., 1994; Yun et al., 2001). It has been repeatedly reported to inhibit

dynein (D’Souza et al., 2006; Kharkwal et al., 2014) and kinesin motility at a concentration as

low as 100 mM (Scholey, 1993; Dentler et al., 1995). Orthovanadate hence is very likely to

mediate the observed complete arrest of kinesin-1 motility. In consequence, an excess of

β-glycerophosphate alone was used to inhibit dephosphorylation of proteins within whole cell

lysates by serving as an alternative substrate. These high concentrations of

β-glycerophosphate, however, interfered with microtubule-motor binding and led to detachment

of microtubules (Figure 8D). Most likely, negatively charged phosphate groups of

β-glycerophosphate (Kolawole et al., 2019) disturb the electrostatic interaction between the

lattice-exposed acidic C-termini of tubulin subunits (E-hook) and the positively charged K-loop of

kinesin-1 (Okada and Hirokawa, 2000). These E-hook/K-loop interactions are crucial to stabilize

the weak binding state of kinesin motor domains (Okada and Hirokawa, 2000), resulting in more

frequent detachment of microtubules from surface-bound kinesin-1 in the presence of

β-glycerophosphate. To prevent this, methylcellulose was added to all solutions in the gliding

assay, which increases the viscosity of the assay solution. This prevents detached microtubules



from escaping into solution, keeps them closer to the imaging surface and facilitates their

efficient recapture by surface-bound motors (Wiesner et al., 2003; Saito et al., 2017; Farhadi et

al., 2018).

Implementation of these adjustments in the used kinesin-1-dependent microtubule gliding

assay produced highly reproducible results (Figure 9D). In line with previous studies,

kinesin-1-dependent microtubule gliding was completely inhibited at concentrations of

recombinant human 2N4R tau-GFP of around 118 nM and higher (Figure 10C) (Hagiwara et al.,

1994; Peck et al., 2011; Parimalam et al., 2016). The same sensitivity still applies to the used

kinesin-1-dependent microtubule gliding assay in the presence of whole cell lysates, underlining

the robustness and suitability of the modified assay to study the influence of proteins in such a

protein-rich solution on kinesin-1 motility.

Comparison of chemical and mechanical protein isolation techniques

Various protein isolation techniques were utilized in this study to determine the method yielding

the most sufficient cell lysis in combination with a high concentration of total protein in the final

lysate, while being compatible with the herein used kinesin-1-dependent microtubule gliding

assay. Buffers used for chemical cell lysis yielded the highest amounts of total protein and

sufficiently lysed cells so that the housekeeping protein β-actin could be detected (Table 9).

However, those buffers interfered with microtubule gliding on kinesin-1 motors, leading to

microtubule detachment even in the presence of very highly diluted buffer or after buffer

exchange (Figure 8). Hence, different mechanical lysis methods, utilizing PBS as a buffer that is

not interfering with kinesin-1-dependent microtubule gliding (Figure 8), were tested for their lysis

efficiency and protein yield.

In this study, the housekeeping protein β-actin could not be detected in motor neuron

lysates produced by glass bead shearing (Table 9), suggesting that lysis was incomplete or that

a major portion of protein got lost during lysate processing. Evidence for the latter comes from

the fact that beads are generally provided to the cell suspension and mixed at high speeds to

break open the cell membrane by shear force. This process generates heat, which may cause

degradation of proteins to some degree (Shehadul Islam et al., 2017) and cause the observed

loss of protein. This is in line with the finding that protein denaturation and aggregation has been

observed in yeast cell lysates produced by glass bead shearing (Papanayotou et al., 2010).

Likewise, β-actin could not be detected in lysates prepared by grinding of motor neurons in the

presence of liquid nitrogen (Table 9). Although grinding in liquid nitrogen has been successfully

used in previous studies, e.g. for the isolation of protein from Chlorella vulgaris (Zheng et al.,



2011), the procedure presented several disadvantages in this study. For instance, slight thawing

of the lysate could not be avoided during collection of the finely ground powder from the mortar,

which might have caused some protein degradation. Also, a full recovery of the lysates from the

mortar was not possible. A similar problem arose when preparing cell lysates using a dounce

homogenizer. Consistent with previously reported efficient use of dounce homogenization

(Osborne and Neuhoff, 1972; Gozes et al., 1977; Krishnaswami et al., 2016), dounce

homogenization of single motor neurons yielded comparably high amounts of protein as well as

well detectable levels of the housekeeping protein β-actin in this study (Table 9). However, a

small fraction of the lysate always stuck to the walls of the homogenization tube, resulting in a

reduced volume of recovered lysate. Therefore, mechanical lysis of motor neurons was chosen

as a suitable method for whole cell lysate production in this study, by passing the cell

suspension several times through a thin needle with the help of a syringe. This method resulted

in similar protein yields and detection of β-actin in western blot as compared to dounce

homogenization (Table 9), but nearly full lysate volume recovery since the lysate could be

pressed out of the syringe and needle using the plunger.

4.2 Wildtype and ALS-associated FUS variants do not directly interfere with

kinesin-1 motility and do not bind to microtubules

Defects in the axonal transport of a variety of organelles and nucleic acids have long been

observed in ALS (Fujii and Takumi, 2005; Bilsland et al., 2010; Ikenaka et al., 2012; De Vos and

Hafezparast, 2017b; Naumann et al., 2018; Pal et al., 2018; Smith et al., 2019), the underlying

mechanism for impaired transport is, however, still enigmatic. In this study, a modified

kinesin-1-dependent microtubule gliding assay was used to investigate a direct influence of

ALS-associated FUS variants alone or in combination with their endogenous interaction

partners present in whole cell lysates on the kinesin-1-driven transport machinery. Results

presented herein provide clear evidence that neither recombinant human wildtype FUS-GFP nor

FUS-P525L-GFP by itself directly binds to microtubules or inhibits kinesin-1 motility over a wide

range of concentrations. Although relative microtubule gliding velocity in the presence of FUS

variants slightly reduced over the course of three hours (Figure 12A), this phenomenon can be

likely attributed to aging of the assay components. Aging occurs as a result of changes in pH,

depletion of ATP, or oxidative damage due to depletion of oxygen scavenging components

glucose oxidase, catalase or DTT (Rasnik et al., 2006; Aitken et al., 2008; Landry et al., 2009;

Swoboda et al., 2012). Interestingly, although the median gliding velocity was comparable in the



presence of recombinant FUS variants over the course of three hours and in the presence of

low amounts (15-25 nM) of 2N4R tau-GFP (Figure 12A and Figure 10C), aging of the assay

resulted in a uniform decrease in the gliding velocity of all microtubules gliding in the field of

view, while the introduction of tau-GFP caused only individual microtubules to slow down or fully

stop (Figure 15B and D). The fraction of non-gliding microtubules sharply increased after a

threshold concentration of 15 nM tau-GFP until virtually all microtubules have stopped gliding at

a concentration of 50 nM 2N4R tau-GFP or above. A very similar distribution of gliding and

non-gliding microtubules has been observed in a system employing active and inactive

kinesin-1 motors at different ratios. When 30% of kinesin-1 motors immobilized on a glass

surface were inactive, microtubules displayed a bistable movement with one portion of

microtubules gliding at a peak velocity around 500 nm/s and another portion with peak velocity

at 0 nm/s. Slightly higher or lower amounts of inactive motors shifted the distribution of

microtubule gliding velocities towards the slower or the faster peak, respectively (Scharrel et al.,

2014). Whether a microtubule is propelled by the underlying motors therefore depends on the

number of inactive motors that attach to the filament, where a stop of motion is achieved when

inactive motors dominate. Similarly, whether a microtubule is propelled by underlying kinesin-1

motors in the presence of 2N4R tau-GFP depends on the amount of tau molecules creating

roadblocks on the microtubule, thereby impairing kinesin-1 motility. Since neither FUS variant

tested in the present work caused a bistable kinesin-1-dependent gliding of microtubules, this

suggests that wildtype FUS and FUS-P525L do not impair kinesin-1 motility on microtubules by

direct interaction with either microtubules or kinesin-1 motors.

FUS by itself hence does not interfere with the kinesin-1-driven transport machinery.

Further, whole cell lysates of iPSC-derived spinal motor neurons expressing the same FUS

variants did not interfere with kinesin-1-driven microtubule gliding at the tested concentrations.

This indicates that FUS variants did not form complexes with other proteins in whole cell lysates

that would interfere with kinesin-1 motility. Taken together, these results strongly suggest that

neither the wildtype form nor the mislocalized cytoplasmic FUS-P525L variant directly or by

interaction with other proteins interfere with kinesin-1-driven (anterograde) axonal transport in

neurons.

Yet, there are some limitations to the herein used assay setup which have to be taken into

consideration. Although there were no aggregates of recombinant or endogenous FUS variants

apparent in the presented experiments, even over the course of three hours, ALS-associated

FUS aggregation has repeatedly been observed in vivo and in vitro (Shelkovnikova et al., 2014;

Murakami et al., 2015; Patel et al., 2015; Marrone et al., 2019) and may very well be involved in



hindering axonal transport in neurons of ALS-patients, e.g. by sterical hindrance (Sau et al.,

2011) or sequestering of proteins (Aulas and Vande Velde, 2015; Yasuda et al., 2017). In case

aggregates composed of cytoplasmic mislocalized FUS-P525L-GFP were initially present in

iPSC-derived spinal motor neurons, they were most likely lost during cell lysate clearance by

centrifugation (>100K g in pellet, Shelkovnikova et al., 2014) and hence no longer present in

whole cell lysates. A possible impact of endogenous cytoplasmic FUS aggregates on axonal

transport is difficult to study in a minimal system such as microtubule gliding assays, since

compartmentalization of the neuron is lost during lysates generation. Furthermore, a loss of

compartmentalization after cell lysis also abolishes the possibility to study pathological events

with local impact. For instance, axonal transport of mitochondria is significantly impaired and

their membrane potential broken down in iPSC-derived spinal motor neurons expressing

FUS-P525L-GFP (Naumann et al., 2018), indicating a potential local depletion of ATP and

hence impaired kinesin-1 motility. Local ATP deficiencies, however, are lost during lysis, and an

excess of ATP is additionally supplied to cell lysates used in kinesin-1-dependent microtubule

gliding assays to ensure efficient kinesin-1-motility. Another challenge for studying the impact of

cytosolic inclusions in microtubule gliding assays presents the generation of FUS-containing

aggregates. In vitro assembled FUS aggregates do not form uniformly in size and do not fully

resemble all components present in endogenous pathological FUS aggregates (Marrone et al.,

2019). Endogenous FUS granules could be enriched by centrifugation in fractions from cell

lysates (Shelkovnikova et al., 2014) in the past, however, this required the introduction of the

detergent Triton X to lysates, which has been shown to influence kinesin-1-dependent

microtubule gliding velocity and causes detachment of actin filaments from myosin motors

(Kellermayer, 1997; Kuramitsu, 1999). Endogenous cytoplasmic FUS aggregates are also likely

to sequester proteins and RNA interacting with cytoplasmic FUS under physiological conditions,

which results in severe reduction in their concentration or virtual exclusion from herein used

whole cell lysates during clearance by centrifugation or because the neuronal cytoplasm

becomes highly diluted after lysis. Lack of such factors might hence contribute to the absence of

a potential interference of FUS with kinesin-1-dependent microtubule gliding in this study.

Finally, some interactions of proteins in cell lysates with kinesin-1 might have been missed due

to the inaccessibility of motors in a gliding assay.

Gliding assays present a useful tool to study the effect that proteins have on the transport

machinery, in particularly if they interact with microtubules or the motor domain or motor

proteins, and imaging of large polymeric filaments is not technically challenging. Yet, the actual

imaging of single motors walking across cytoskeletal filaments in stepping assays presents a



more direct way of measuring motor characteristics and how these may change in the presence

of proteins engaging with the motor. Depending on the motor construct that is used (i.e.

full-length kinesin-1) in a stepping assay, the motor itself becomes much more accessible for

interaction with proteins in solution, compared to its surface-bound state in a gliding assay.

Proteins present within the herein used whole cell lysates may have been prevented from

interacting with kinesin-1 due to the inaccessibility of the surface-bound motor. Hence, applying

whole cell lysates of isogenic iPSC-derived spinal motor neurons carrying different FUS variants

in a stepping assay may yield different results than those presented here and provide further

insights into how mutations in FUS impact the kinesin-1-driven transport machinery directly.

In addition, there is evidence that FUS associates with ATP-dependent actin-binding

motor Myo5A (Yoshimura et al., 2006) and Myo6 (Takarada et al., 2009), and has been isolated

as part of a large granule associating with KIF5B (Kanai et al., 2004). Although the herein

presented results demonstrate that FUS by itself does not interfere with kinesin-1 (KIF5B)

motility, it remains to be elucidated whether this holds true for actin-associated myosin motors

as well. Similarly to the kinesin-1-dependent microtubule gliding assay used in this study, actin

filaments can glide over myosin-coated surfaces (Rock et al., 2000; Vilfan, 2009; Hariadi et al.,

2016), and single myosin molecules can be tracked while stepping on immobilized actin

filaments (Ricca and Rock, 2010; Bao et al., 2013). By utilizing such actin and myosin based

gliding or stepping assays in future experiments, the direct interaction of FUS variants with

these components of the transport machinery may be investigated.

Taken together, the present data provide strong evidence that mislocalized, free

cytoplasmic FUS-P525L itself does not directly interfere with kinesin-1-driven axonal transport

on microtubules. Apart from the fact that ALS-associated FUS aggregates sequester proteins

involved in RNA transport (Kamelgarn et al., 2016) as well as mRNA and protein of kinesin-1

itself (Yasuda et al., 2017), there is also little evidence in the literature to date about a direct

involvement of mislocalized cytoplasmic ALS-associated FUS in microtubule-based axonal

transport.

4.3 An increase in 4R:3R tau isoform ratio might contribute to the axonal

transport defects observed in FUS-ALS

Since the herein presented results show that FUS is not directly interfering with the microtubule

based kinesin-1 mediated transport machinery, this study also investigated possible effects of

FUS-dependent changes in the expression of tau isoforms on axonal transport. Defects in



axonal transport have been observed early in the progression of many neurodegenerative

diseases, and might partly be caused by hyperphosphorylated mutant tau variants or by altered

expression levels of tau isoforms (Ebneth et al., 1998; Stamer et al., 2002; De Vos et al., 2008;

Goldstein, 2012; Qian and Liu, 2014; Correia et al., 2016; Fernández-Nogales and Lucas,

2020). Overexpression of single wildtype tau isoforms (0N3R, 2N3R, 0N4R, and 2N4R) in H4

human neuroglioma cells have previously been associated with defects in axonal transport of

mitochondria (Stoothoff et al., 2009). Previous studies could show that the FUS, together with

the transcription factor SFPQ, is directly involved in the splicing of tau pre mRNA, and a

knockdown of FUS results in in a higher expression of the 4R tau isoform in mice (Ishigaki et al.,

2017; Bourefis et al., 2020). Furthermore, pathological behavioral phenotypes caused by FUS

knockdown in these mice could be weakened by silencing of 4R tau with small hairpin RNAs

(Ishigaki and Sobue, 2018), indicating that an increased 4R:3R ratio within the cell may indeed

contribute to pathology. In line with this, changes in the 4R:3R in favor of 4R have been

observed in diverse tauopathies (D’Souza and Schellenberg, 2005; Goedert and Jakes, 2005;

Sergeant et al., 2005). However, the role of changing ratios of wildtype 4R and 3R tau isoforms,

without increased total tau levels, in contributing to axonal transport defects is not well

investigated to date.

We hence hypothesized that cytoplasmic mislocalization of FUS and the resulting loss of

function in splicing of tau might as well increase the 4R:3R isoform ratio of tau in vivo, which in

turn might affect kinesin-1-dependdent transport in axons. In order to test this hypothesis, the

microtubule binding affinity of purified recombinant human 2N3R tau-GFP and

2N4R tau-mScarlet and their effect on kinesin-1-dependent microtubule gliding was analyzed in

the modified microtubule gliding assay.

Although the largest tau isoforms utilized in this study are the least expressed within the

human brain (~9%, Hong, 1998), the ratio between respective 3R and 4R isoforms with varying

N-termini (e.g. 0N3R and 0N4R) is similar and hence results obtained with 2N3R and

2N4R tau isoforms are expected to be transferable to all other tau isoforms (He et al., 2020).

While both tau variants used in this study impaired microtubule gliding in the modified

kinesin-1-dependent microtubule gliding assay, a 5-fold lower concentration of

2N4R tau-mScarlet than of 2N3R tau-GFP variant was sufficient to slow relative microtubule

gliding speed down to 0.19 ± 0.09 and 0.31 ± 0.13, respectively (Figure 17C). In addition, an

approximately 20-fold higher microtubule binding affinity was measured for 2N4R tau-mScarlet

(Kd = 6.79 ± 0.41 nM) compared to 2N3R tau-GFP (Kd = 137.12 ± 29.7 nM) (Figure 18C, D). In

both cases the binding data was best fitted with a Hill-model fit with a Hill coefficient of



2.38 ± 0.67 for 3R tau GFP and 1.83 ± 0.29 for 4R tau mScarlet (Figure 18C). Both coefficients

are larger than 1, suggesting a cooperative binding mechanism for 2N3R and 2N4R tau

isoforms (Weiss, 1997). This means that the binding of tau molecules to microtubules may not

be independent, but that binding of one tau molecule increases the binding-probability of

another tau molecule. While the exact mechanism by which tau binds soluble tubulin or

microtubules is still elusive, Fung et al. have proposed a model for its cooperative binding to

tubulin dimers (Fung et al., 2020), and other recent studies also suggest a cooperative binding

mechanism of 2N4R tau within non diffusive, but dynamic tau condensates on microtubules

(Siahaan et al., 2019; Tan et al., 2019).

Previous studies often used co-sedimentation assays to determine the binding affinity of

3R and 4R tau isoforms to microtubules, and results varied greatly from up to 6 µM down to less

than 100 nM (Butner and Kirschner, 1991; Goode and Feinstein, 1994; Hong, 1998; Ackmann et

al., 2000; Goode et al., 2000). However, bound tau was in those cases assessed by SDS-PAGE

gel analysis, which is not suitable to reliably detect tau concentrations below 1 µM (Gustke et

al., 1994b). And yet, studies performing a much more sensitive fluorescence resonance energy

transfer (FRET) assay or nuclear magnetic resonance (NMR) also reported largely varying

dissociation constants for comparable tau isoforms, ranging from 2.5 µM down to 14 nM

(Makrides et al., 2004; Sillen et al., 2007; Elbaum-Garfinkle et al., 2014; Maïo et al., 2014). Also,

in contrast to these studies, microtubule bundles were excluded from the analysis in this study.

The quantification of dissociation constants for tau hence largely depends on the experimental

procedures and can be compared only to a very limited extent.

However, while the exact Kd-values may vary, these studies consistently reported a

stronger microtubule binding affinity for 4R compared to 3R tau variants, which is consistent

with the herein presented results. This suggests that when the ratio of 4R:3R tau isoforms

changes, the microtubule decoration with tau isoforms and hence the motility of kinesin-1

walking across these microtubules may change as well. To test this hypothesis, we analyzed

the relative microtubule-binding behavior of mixed recombinant human 4R tau-mScarlet to

3R tau-GFP and their isoform-ratio dependent effect on kinesin-1-dependent microtubule

gliding. Increasing 4R:3R tau isoform ratios led to decreased relative microtubule gliding

velocities (Figure 19), suggesting that a shift towards higher 4R tau isoform levels within a

neuron negatively affects axonal transport along microtubules. Interestingly, the fact that

2N4R tau-mScarlet in solution seems to prevent 3R tau-GFP from binding to microtubules, even

at low 4R:3R ratios (Figure 20), suggests that transport defects are caused by a higher

decoration of axonal microtubules with 4R tau isoforms due to its higher roadblock efficiency.



Together, these results suggest that an increased 4R:3R tau isoform ratio within neurons

expressing the FUS-P525L variant likely contributes to, or might even be sufficient to cause the

axonal transport defects observed in FUS-ALS pathology.

Due to the higher net positive charge of the microtubule binding domain, 4R tau also

shields the negative surface charge of microtubules more efficiently when compared to 3R tau.

This results in a reduced kinesin-1 landing rate and decreased kinein-1 velocities (Parimalam et

al., 2016). In line with these findings in vitro, 0N4R tau largely co-localized with microtubules of

NIH 3T3 cells when co-transfected with differently labeled 0N3R and 0N4R tau isoforms at a 1:1

ratio, while 0N3R tau diffusely distributed to the cytosol (Lu and Kosik, 2001). Studying the

effect of 4R:3R tau isoform imbalances in mouse models is difficult, as 4R tau is the

predominantly expressed isoform in the adult rodent brain while 3R tau is hardly expressed

(Lacovich et al., 2017; Tuerde et al., 2018). Therefore, Lacovich et al. created a trans-splicing

model in neurons differentiated from human embryonic stem cells (hESCs), which allowed for

selective in- and exclusion of exon 10 leading to an increase in relative endogenous levels of 3R

or 4R tau isoforms, respectively. Using this model system, they could show that an increased

level of 3R tau isoforms relative to 4R tau favored anterograde transport of the amyloid

precursor protein (APP), while an increased 4R:3R tau isoform ratio promoted

(dynein-dependent) retrograde transport by decreasing (kinesin-dependent) anterograde

transport velocities (Lacovich et al., 2017). Although these neurons differentiated from hESCs

predominantly express 3R tau isoforms, this data suggests that a shift in 4R:3R tau isoform ratio

is sufficient to interfere with the transport of APP in vivo. Taken together, previous studies in line

with the data presented herein provide clear evidence that a slight shift in 4R:3R tau isoform

ratio, e.g. caused by the nuclear loss of function of FUS-P525L, impairs kinesin-1-dependent

transport on microtubules.

Interference of other tau isoforms with kinesin-1 motility

Although the effect of tau on motor motility has been studied in detail, studies often investigated

only one tau isoform or variant, preventing any direct cross-comparison of tau isoforms and their

ability to interfere with kinesin-1 motility on microtubules. In addition, taxol-dependent

microtubule stabilization or the decoration of microtubules with tau differentially affects kinesin-1

motility. For instance, microtubules assembled in the presence of 0N3R tau glide moderately,

but significantly faster (0.54 ± 0.01 µm/s) compared to microtubules assembled in the presence

of 0N4R tau (0.48 ± 0.01 µm/s) (Yu et al., 2014). Similarly, another study reported a significantly

faster gliding of microtubules assembled in the presence of 0N3R tau (0.52 µm/s) than in the



presence of 0N4R tau (0.43 µm/s) (Peck et al., 2011). When microtubules are, however,

assembled in the presence of taxol and subsequently decorated with these tau isoforms, the

gliding velocity is slower in the presence of 0N3R tau (0.29 ± 0.01 µm/s) compared to 0N4R tau

(0.32 ± 0.01 µm/s) at tau concentrations of 100-1000 nM (Yu et al., 2014).

The properties of molecular motors are also often studied in so called stepping assays,

where either single motor molecules or cargo-attached motors are allowed to bind to and

translocate along surface-immobilized microtubules, while they are visualized by TIRF

microscopy. These assays are more suitable to investigate the characteristics of single motors

when encountering proteins interfering with motor motility on microtubules compared to

microtubule gliding assays, where motors act in ensemble to propel microtubules across the

surface. In single molecule stepping assays, kinesin-1 moves at similar velocities in the

presence of 0N3R tau (0.36 ± 0.02 µm/s) or 0N4R tau (0.33 ± 0.02 µm/s), but shows a

decreased run length at these conditions (Vershinin et al., 2007; Dixit et al., 2008; Yu et al.,

2014). Conversely, concentrations of 2N4R tau up to 1.5 µM did not decrease the run length of

recombinant rat kinesin-1, but drastically reduced its landing on microtubules, leading to the

hypothesis that the crucial step in the motility of this kinesin-1 is its attachment to microtubules,

where it translocates undisturbed by tau roadblocks once it is bound (Seitz et al., 2002). A

similar effect of 2N4R tau has been observed in CHO cells. The velocities of a variety of cellular

organelles and vesicles were unaffected upon microinjection of recombinant 2N4R tau into

these cells, but attachment and detachment frequencies were altered causing impaired

transport of these organelles and vesicles throughout the cell (Trinczek et al., 1999).

Interestingly, low concentrations of 2N4R tau (0.25-20 nM) lead to the reversible formation of

cohesive islands or condensates on taxol-stabilized (but not GMP-CPP-stabilized) microtubules

(Siahaan et al., 2019; Tan et al., 2019). These condensates seem to have a kind of regulatory

effect on motor translocation across microtubules, as they are constitute impermeable barriers

for kinesin-1 motors but not for processive dynein-dynactin-BICD2N (DDB) complexes, while

they are disassembled by kinesin-8. Tau islands further protect microtubules from severing

enzymes such as katanin (Siahaan et al., 2019) or spastin (Tan et al., 2019). In vitro studies

suggest that alternative splicing of tau does not seem to affect tau condensate formation on

microtubules, since 2N3R and 0N3R tau readily incorporated into preformed 2N4R tau

condensates (Tan et al., 2019). Further, it has been hypothesized that impaired motor motility

on microtubules upon higher 4R levels in solution is not caused by an overall higher decoration

of microtubules with tau molecules, but due to the replacement of tau molecules with three



microtubule binding repeats by those with four repeats, as it has been observed in vivo (Lu and

Kosik, 2001).

Interestingly, the presence of increasing concentrations (1-100 nM) of the longest tau

isoform (2N4R) caused a more subtle decrease the run length of unloaded, GFP-labeled kinesin

in stepping assays, compared to the presence of the shortest isoform (0N3R) (Vershinin et al.,

2007; Dixit et al., 2008), indicating that the projection domain of tau may actually counteract the

roadblock effect caused by tau binding to microtubules and aid in kinesin translocation. In line

with this, Tarhan et al. have investigated all six human tau isoforms for their capacity to act as

roadblocks for bead-attached kinesin-1 motors. Increasing concentrations of 2N4R tau

(0.14 - 140 nM) resulted in a significant decrease of bead velocity and hence kinesin-1

translocation speed. While all 4R tau isoforms had a higher microtubule binding affinity and

decreased kinesin-1 velocity along microtubules stronger compared to their respective 3R tau

N-terminal counterparts (e.g. 0N3R compared to 0N4R), the kinesin-1 velocity increased for tau

isoforms with more N-terminal domains (0N<1N<2N). This suggests that the negatively

charged projection domain of tau aids in recruiting and keeping positively charged kinesin-1

motor domains to the microtubule lattice (Tarhan et al., 2013), and in combination with a

previously proposed sidestepping mechanism allowing kinesin to bypass roadblocks on

microtubules (Dixit et al., 2008; Yildiz et al., 2008; Dreblow et al., 2010). Intriguingly, an

additional N-terminal domain (i.e. in 1N3R tau) was reported to cause a considerably larger

slowdown in microtubule gliding velocity compared to 0N3R tau (on taxol-stabilized

microtubules decorated with 20, 200, or 400 nM of the respective recombinant tau isoform)

(Schmidt et al., 2012).

In light of these sometimes opposing results in previous studies, it would be interesting to

investigate the interference of the different tau isoforms, alone or in combination with each

other, utilized in this present study with individual kinesin-1 motors in the reversed assay

geometry of stepping assays. In addition, the fact that the projection domain of tau may

influence the stepping behavior of kinesin-1 on microtubules suggests that stepping assay in the

presence of different ratios of all six 3R and 4R isoforms would add valuable knowledge as to

how a shift in 4R:3R tau isoform ratio could impair kinesin-1 motility on microtubules within the

cell.

Batch-to-batch differences in purified recombinant 2N4R tau variants

Initial experiments of this study were performed with recombinant human 2N4R tau-GFP, kindly

provided by Dr. Marcus Braun from the Institute of Biotechnology CAS in Prague, while later



experiments were performed with 2N4R tau-mScarlet, purified by myself. Both batches exhibit

significantly different inhibitory potential in the kinesin-1-dependent microtubule gliding assay.

While kinesin-1-dependent microtubule gliding completely ceased in the presence of 118 nM of

the 2N4R tau-GFP variant (Figure 10), about 10-fold more protein (1200 nM) of the

2N4R tau-mScarlet variant was needed to reduce the relative microtubule gliding velocity to

0.18 ± 0.09 (Figure 17). In order to determine to what degree different batches of tau differ in

their inhibitory potential and to exclude variations due to the different fluorophores, two more

2N4R tau batches, i.e. 2N4R tau-mCherry and 2N4R tau-GFP, were purified and tested in the

modified kinesin-1-dependent microtubule gliding assay (Figure S2). Significantly larger

amounts of 2N4R tau-mScarlet (1200 nM), -mCherry (1500 nM), or –GFP (1500 nM, second

batch) were required to reduce microtubule gliding velocity down to 0.18 ± 0.09, 0.15 ± 0.01, or

0.44 ± 0.01, respectively. However, the qualitative effect on kinesin-1 motility was the same for

all tested tau isoform variants, since the addition of increasing amounts of tau consistently

decreased kinesin-1-dependent microtubule gliding velocity. Together with above discussed

previous studies, where the concentration of tau used ranges from nM to mM, it indicates that

the inhibitory potential of tau varies between different purification batches (Trinczek et al., 1999;

Seitz et al., 2002; Vershinin et al., 2007; Dixit et al., 2008; Peck et al., 2011; Schmidt et al.,

2012; Tarhan et al., 2013; Yu et al., 2014; Siahaan et al., 2019; Tan et al., 2019).

The absolute amount of a protein needed to reproduce an effect or degree of activity often

differs between individual batches of purified recombinant protein. During the purification of the

protein of interest, co-purification of endogenous proteins (in this case from insect cells)

commonly occurs, while the composition and amount of these co-purified proteins differs from

batch to batch (Structural Genomics Consortium et al., 2008). Such proteins could include

MAPs or motors, which are unlabeled and hence invisible for widefield-epifluorescence, but

would interfere with kinesin-1 motility in a microtubule gliding assay. Further, intrinsically

disordered proteins like tau (KrishnaKumar and Gupta, 2017) tend to form aggregates, a

process often aggravated by long term storage of proteins (Chakraborty et al., 2018).

Aggregated tau likely impairs kinesin-1 motility on microtubules differently compared to

non-aggregated, soluble tau. In addition, the activity of tau strongly depends on its

phosphorylation state. The microtubule binding affinity of tau is decreased upon its

hyperphosphorylation (Ballatore et al., 2012). When purified from insect cell, tau is heavily

phosphorylated (Tepper et al., 2014), although the exact degree of phosphorylation is not

consistent between individual purifications. Hence, different phosphorylation states between the



earlier purified 2N4R tau-GFP and the later purified 2N4R tau-GFP or -mScarlet, which likely

influence its ability to hinder kinesin-1 motility on microtubules, cannot be excluded.

The up- and downsides of using Taxol in combination with tau

The modified kinesin-1-dependent microtubule gliding assay presented in this study used in

vitro polymerized microtubules that were stabilized by the addition of taxol. Previous studies

showed that taxol can migrate through 2-nm pores within the microtubule wall, where it binds to

the luminal face of β-tubulin (Meurer-Grob et al., 2001; Li et al., 2002; Kar et al., 2003; Ross and

Fygenson, 2003) and counteracts the effect of GTP hydrolysis (i.e. tubulin destabilization) on

the outward facing side of this β-tubulin molecule (Amos and Löwe, 1999; Prota et al., 2013).

Interestingly, recent reports suggest that tau molecules occupy a similar, if not the same binding

site in the microtubule lumen as taxol (Kar et al., 2003; Ross et al., 2004; Yu et al., 2014), with

the N-terminus of tau projecting outside the microtubule wall (Amos, 2004a, 2004b), a process

which primarily occurs if microtubules are assembled in the presence of tau and is thought to be

irreversible (Makrides et al., 2004). This indicates that, because microtubules in the herein used

assay were stabilized with taxol before tau addition, some binding sites for tau on the

microtubules were likely already occupied by taxol molecules, which would result in less tau

molecules interacting with the microtubule lattice. However, the incorporation of tau into the

microtubule lumen is expected to be especially important for the stabilization of dynamic

microtubules, and only to a lesser extent for the interaction with other MAPs such as motors. In

addition, while tau was found to induce cooperative binding of Taxol to microtubules, the inverse

was not observed as binding of tau to microtubules was independent of the presence or

absence of Taxol during microtubule assembly (Ross et al., 2004). Tau has also been found to

reversibly bind to the outer surface of microtubules (Makrides et al., 2004; Martinho et al.,

2018), along as well as across filaments (Santarella et al., 2004; Duan et al., 2017), a process

more likely to influence microtubule-dependent transport. Hence, the stabilization of

microtubules with taxol before tau addition, as performed in the herein presented experiments,

is considered to have only minor impact on the herein measured microtubule gliding velocities.

Tau isoform expression profile in iPSC-derived neuronal cultures

Intriguingly, whole cell lysates obtained from patient-specific iPSC-derived spinal motor neurons

expressing either the wildtype FUS-GFP or the FUS-P525L-GFP variant did not differentially

interfere with kinesin-1-dependent microtubule gliding (Figure 13B). Due to the cytoplasmic

mislocalization of FUS-P525L-GFP, the 4R:3R tau isoform ratio is expected to be increased in

lysates from cells expressing this variant. Hence we would expect the relative microtubule



gliding velocity to decrease in lysates from neurons expressing FUS-P525L-GFP compared to

lysates from cells expressing wildtype FUS-GFP. Therefore, the level of total 3R and 4R tau

isoforms in the used whole cell lysates was determined by western blot using 3R or 4R isoform

specific antibodies, respectively (Figure 21).

However, only the shortest, 0N3R isoform could be detected in cell lysates irrespective of

the expressed FUS variant, indicating that all other isoforms are not expressed above the

detection limit of this technique (low pg range, Wiltfang et al., 1991). This is in line with other

studies on iPSC-derived neurons (Iovino et al., 2015; Verheyen et al., 2015, 2018; Biswas et al.,

2016; Imamura et al., 2016; Silva et al., 2016; Hallmann et al., 2017; Sato et al., 2018) and

reflects the relative immaturity of these reprogrammed neurons, since 0N3R tau is the only

isoform expressed in the fetal human brain (Kosik et al., 1989; Goedert and Jakes, 1990). While

other tau isoforms can be detected on the RNA level in iPSC-derived neurons (Ehrlich et al.,

2015), detection of the 0N4R, 1N3R, and 1N4R isoforms on the protein level is only possible

when neurons are allowed to mature for up to 365 days (Iovino et al., 2015; Sposito et al.,

2015). In fact, recent evidence suggests that iPSC-derived neurons display transcriptional

profiles similar to those found in human fetal brains rather than adult brains (Handel et al., 2016;

Sobol et al., 2019; Griesi-Oliveira et al., 2020).

Taken together, we have to assume that the whole cell lysates used in this study, obtained

from iPSC-derived spinal motor neurons at day 21 of maturation, did not contain endogenous

tau isoforms other than the shortest, fetal 0N3R isoform due to the relative immaturity of

differentiated neurons. Although, previous studies reported defects in axonal transport in

mitochondria in the iPSC-derived spinal motor neurons as early as day 21 of maturation

(Naumann et al., 2018). This implies that the axonal transport defects observed in iPSC-derived

spinal motor neurons are not caused by a shift in tau isoform splicing. However, further

evidence on the transcriptional and protein level would be needed to consolidate this

hypothesis.

In order to obtain more mature neurons, the time that neurons are allowed to differentiate

in culture could be increased, while the course of tau isoform expression levels within these

neurons after defined time points should be evaluated. Other studies have demonstrated the

culture of iPSC-derived spinal motor neurons up to almost 130 days (Devlin et al., 2015), when

the fetal 0N3R tau isoform is however still predominantly expressed (Iovino et al., 2015; Sposito

et al., 2015). Unfortunately, with increasing culture time, iPSC-derived spinal motor neurons

tend to form clusters over time and detach from the herein used PLO-laminin coated surface.



Another recently evolved method to obtain disease- or patient-specific neurons, and to

circumvent the rejuvenation or reset in maturation state during the production of iPSCs, is direct

programming of neuronal subtypes from human somatic cells. A number of somatic cell types,

including mouse and human fibroblasts, but also astrocyte and human retina-derived fibroblasts,

can be transformed into a variety of neuronal subtypes using different combinations of

overexpressed transcription factors, small molecules or the application of microRNAs (reviewed

in Liou et al., 2020).

In addition, cells cultured in a 3D culture system have recently proven to relate to the

characteristics, including transcriptional and expression profiles, of their in vivo counterparts

more closely compared to cells grown in conventional 2D culture systems (Kapałczyńska et al.,

2016; Duval et al., 2017). Especially the reconstruction of retinal development and regeneration

has been extensively studied using organoids created in a 3D culture of human iPSCs (Eiraku

et al., 2011; Zhong et al., 2014; Hallam et al., 2018; Shrestha et al., 2019). However, 3D culture

systems have also been used to model AD and have enhanced the maturation of iPSC-derived

cortical neurons. Interestingly, neurons grown in this 3D culture exhibit an increased RNA

expression level of 4R tau isoforms already after 49 days in culture (Choi et al., 2014). A

systemic analysis of tau isoform protein expression levels in a 3D culture system revealed an

expression of 0N3R, 0N4R, 1N3R and 1N4R tau isoforms after 15 weeks in culture (105 days),

while 2N3R and 2N4R tau isoforms were expressed after 25 weeks in culture (175 days). As in

the 2D culture system though, the 0N3R tau isoform remained the predominantly expressed

isoform throughout all time points (Miguel et al., 2019).

Although the generation of mature spinal motor neurons can hence be improved through

the above discussed culture conditions, the tau isoform expression profile within these human

iPSC-derived neuronal subtypes presents the major limitation for studying the interference of an

increased 4R:3R tau isoform ratio on kinesin-1-dependent axonal transport. However, the fact

that defects in axonal transport have been observed in iPSC-derived neurons at early stages of

maturation indicates that those defects must be due to another, non-tau related pathogenesis.

4.4 FUS variants may indirectly affect microtubule-based axonal transport

by acting on a multitude of cellular processes

As FUS affects multiple cellular pathways, cytoplasmic mislocalized ALS-associated FUS may

also indirectly interfere with microtubule-based axonal transport. Due to its diverse roles in DNA

damage repair (Bertrand et al., 1999), RNA metabolism and transcriptional regulation (Yang et



al., 1998), and mRNA transport within the cell (Zinszner et al., 1997), mutant FUS variants can

trigger many, not mutually exclusive, pathological events within the cell, which in turn may

indirectly affect axonal transport.

Nuclear loss of function - DNA damage repair and RNA splicing

Wildtype FUS localizes to sites of laser-induced DNA damage prior to the recruitment of other

key proteins involved in DNA-repair, whereas recruitment of FUS-P525L to sites of

laser-induced DNA damage is impaired in iPSC-derived spinal motor neurons, subsequently

leading to defects in mitochondrial transport (Naumann et al., 2018). Interestingly, the inhibition

of PARG, which degrades the PAR chains attracting FUS to DNA damage (Lin et al., 1997),

rescued not only FUS recruitment to laser-induced DNA damage sites, but also mitochondrial

transport defects, suggesting that impaired DNA damage repair causes defects in axonal

transport (Naumann et al., 2018). In addition, it was shown that DNA damage enhances

cytoplasmic mislocalization of FUS-P525L and induces FUS aggregation as well as distal

axonal trafficking defects (especially of mitochondria) and subsequent distal axonopathy (dying

back of axons) (Higelin et al., 2016; Naumann et al., 2018). This supports the notion that defects

in DNA-repair mechanisms due to cytoplasmic mislocalization of FUS inevitably result in the

long-term accumulation of DNA damage, which coincides with the rather late mean age of onset

in ALS. As neurons lack the ability to self-renew or replicate, they are particularly susceptible

DNA damage accumulating over time which consequently likely causes disruptions in other

cellular pathways, such as axonal transport (Guo et al., 2020).

Next to role in DNA damage repair, FUS is also involved in the expression and splicing of

a variety of motor proteins (Colombrita et al., 2012), such as KIF5B. Accordingly, cytoplasmic

mislocalized mutant FUS variants (i.e. FUS-R521G and FUS-R521H) can no longer partake in

proper splicing of these motors (Hoell et al., 2011). Additionally, FUS binds to mRNAs coding for

β-actin and the actin-stabilizing protein Nd1-L (Fujii and Takumi, 2005). Altered splicing caused

by mutant FUS variants hence likely directly affects cytoskeletal composition, which might affect

axonal transport.

Endosomal trafficking and axonal transport

Clear evidence for how FUS is involved in both, short- and long-distance cellular transport is to

date still sparse. Only few studies have focused on the direct influence of FUS variants on the

axonal transport machinery. One study found that several FUS mutant variants (G230C, R495X,

and R521G) disrupt bidirectional fast axonal transport via activation of the mitogen-activated



protein kinase p38 when expressed in squid axoplasm and in mouse primary motor neurons

(Sama 2017). p38 phosphorylates kinesin-1 and thereby inhibits its translocation along

microtubules, possibly by reducing the landing rate on microtubules (Morfini et al., 2013).

Apart from long-distance transport along the dendritic arbor or axon, short-distance

transport of vesicles, e.g. from the ER to the Golgi apparatus, was also found to be impaired in

FUS-ALS. In particular, Rab-1-dependent transport of coat protein complex II-coated vesicles

from the ER to the Golgi was found to be disrupted upon expression of several FUS-NLS

mutant variants (but not wildtype FUS) (Soo et al., 2015). The same study also showed

inhibition of Golgi-associated vesicular trafficking. Defects in axonal transport of ER-vesicles

was also observed in spinal motor neurons that were differentiated from ALS-patient-derived

iPSCs carrying the FUS-R521H or FUS-P525L variants (Guo et al., 2017b). Interestingly, mice

expressing a frame-shift mutant FUS variant did not show major deficits in endosomal trafficking

(Sleigh et al., 2020), indicating that impaired transport mechanisms in FUS-ALS might not affect

endosomal trafficking in this model.

Mitochondrial damage and trafficking defects

Mutant FUS variants have extensively been linked to the impaired transport of larger organelles,

such as mitochondria. Defective mitochondrial transport may result from the fact that FUS

regulated the expression of mRNAs coding for several motor proteins, such as KIF5C and

KIF1B (Schwarz, 2013; Campbell et al., 2014; Guo et al., 2020). An increase in stationary

mitochondria was observed upon expression of human FUS-P525L, but also wildtype FUS, in

mouse cortical neurons. The same was observed for the fly homologues cabezaP398L and

Cabeza in Drosophila motor neurons (Baldwin et al., 2016; Chen et al., 2016a), suggesting that

overexpression and hence increased cytoplasmic abundance of either FUS variant might

already impair mitochondrial axonal transport. Similarly, the number of motile mitochondria was

significantly decreased in ALS-patient-specific iPSC-derived spinal motor neurons expressing

FUS-R521H or FUS-P525L when compared to isogenic controls (Guo et al., 2017b).

Interestingly , mitochondrial transport defects could be rescued by HDAC6 inhibition, leading to

increased α-tubulin acetylation and hence stabilizing microtubule protofilaments (Janke and

Montagnac, 2017). In addition, an arrest of mitochondria (and lysosome) motility was observed

in the distal axon of ALS-patient specific iPSC-derived spinal motor neurons expressing the

FUS-P525L variant, accompanied by a breakdown in membrane potential and reduced

mitochondrial length (Naumann et al., 2018). ATP production ceases upon the breakdown of

mitochondrial membrane potential, leading to a local depletion of energy around the damaged



mitochondrion (Stoica et al., 2016). A recent study has linked ALS-associated FUS-P525L to

aberrant mitophagy, since the expression of members of the mitochondrial quality control PINK1

and Parkin, as well as ubiquitination of Miro1, is increased in cells expressing this FUS variant

(Park et al., 2006; Sarraf et al., 2013; Chen et al., 2016a, 2016b). This indicates a higher degree

of mitophagy in the presence of FUS-P525L, which also leads to a reduced ATP production.

ATP depletion has been reported to lead to microtubule depolymerization (Park et al., 2013) and

may also locally impair axonal transport due to the lack of the motor’s energy source. Further,

ATP has recently been identified to inhibit fibrillization of pathological FUS aggregates by

specifically binding to its RRM domain (Kang et al., 2019). This implicates that mitochondrial

damage might not only affect axonal transport through ATP depletion, but also by worsening

pathogenic effects of aggregated FUS.

RNP and stress granules

mRNAs assemble with RBPs into ribonucleoprotein (RNP) granules, which are transported

through the axon by active bi-directional transport (Knowles et al., 1996; Tiruchinapalli et al.,

2003; Leung et al., 2006; Nalavadi et al., 2012; Alami et al., 2014; Medioni et al., 2014; Gopal et

al., 2017; Wong et al., 2017; De Graeve and Besse, 2018; Turner-Bridger et al., 2018;

Vijayakumar et al., 2019). Several studies have previously demonstrated that the transport of

RNPs containing FUS depends on the interaction with other RNA transport-related proteins,

such as TDP-43 and SMN (Groen et al., 2013), IMP1 (Kamelgarn et al., 2016), Sam 68 (Belly et

al., 2005), or the adenomatous polyposis coli (APC) tumor-suppressor protein, which targets

β2B-tubulin mRNA to the plus end of detyrosinated microtubules where APC promotes tubulin

synthesis (Mili et al., 2008). Perturbing this interaction leads to disrupted dynamic microtubules

within axonal growth cones (Yasuda et al., 2013; Preitner et al., 2014), and suggests that

structural changes in the cytoskeleton might be a potential cause for axonal transport defects.

Other studies have proposed that RNP granules hitchhike on membrane-bound organelles

when those are transported throughout the cell (Jansen et al., 2014; Salogiannis and Reck-

Peterson, 2017). For instance, recent work has demonstrated a role for kinesin- and

dynein-dependent endosome trafficking in mRNA transport (Gould and Lippincott-Schwartz,

2009; Baumann et al., 2012; Pohlmann et al., 2015). The lysosome has been proposed as

another cellular organelle acting as a platform for RNP granule transport, with coupling possibly

mediated by Annexin A11 (Markmiller et al., 2018; Liao et al., 2019). It will be interesting to

investigate whether mutant FUS variants show perturbed interaction with membrane-bound

organelles and thereby hinder the transport of RNP granules.



After liquid-to-solid phase transition (Murakami et al., 2015; Patel et al., 2015; Bowden

and Dormann, 2016), solidified granules have lost their dynamic properties and can hence no

longer perform their physiological functions (Bowden and Dormann, 2016). The incorporation of

ALS-associated FUS variants into RNP or stress granules and their subsequent pathogenic

phase transition indicates that FUS may impact axonal transport in several distinct, but not

mutually exclusive ways. These include the sequestration of mRNAs and proteins directly

involved in the axonal transport machinery or inhibition of translation of proteins important for

cell survival, such as antiapoptotic factors under prolonged stress conditions (Nevins et al.,

2003). Additionally, when stress granules increase in size (Baron et al., 2013) to form

pathological aggregates, e.g. through the continuous incorporation of RBPs and mRNA, they

might sterically impair cellular processes, i.e. cytoskeletal assembly or axonal transport. A

similar process has been described due to the swelling of mitochondria, which impaired

mitochondrial and lysosomal transport in rat primary neurons (Kaasik et al., 2007).

While it is known that the cytoplasmic localization of FUS on its own is not sufficient to

trigger FUS aggregation or inclusion into RNP and stress granules (Bosco et al., 2010;

Dormann et al., 2010; Ito et al., 2011; Bentmann et al., 2012), there is no clear consensus yet

as to which domain is mainly responsible for the association of FUS with stress granules. Some

evidence points towards its low-complexity, prion-like domain (Sun et al., 2011b; Han et al.,

2012), while others suggest the methylation of arginine residues within the RGG domain RGG

domain(s) (Bentmann et al., 2012; Baron et al., 2013) to be essential for incorporation

(Rappsilber et al., 2003; Dormann et al., 2012). However, its capacity to bind mRNA is clearly

essential for FUS incorporation into granules, as mutations in FUS abolishing mRNA-binding

greatly reduce aggregate-related toxicity (Andersson et al., 2008; Elden et al., 2010; Sun et al.,

2011b; Bentmann et al., 2012; Daigle et al., 2013). This led to the hypothesis that FUS might

undergo conformational changes upon binding to mRNA (Wang et al., 2015; Hamad et al.,

2020), after which it might be capable to interact with an extended set of proteins including

motor proteins or tubulin. It would hence be interesting to see whether microtubule gliding

characteristics would change or whether FUS aggregation occurs in our kinesin-1-dependent

microtubule gliding assay in the presence of sufficient amounts of mRNA transcripts that are

known to interact with FUS.



4.5 Outlook

The present results strongly indicate that FUS does not directly interfere with kinesin-1 or

microtubules. However, it is possible that structurally changed FUS protein or its aggregate

form, e.g. induced by its interaction with mRNAs, may alter its potential to interact with motors or

cytoskeletal filaments. It will therefore be interesting to test whether FUS interferes with

kinesin-1-dependent transport in the presence of mRNA. Further, may not only interfere with

microtubule-associated motors, but also with actin-associated motors, such as myosin. For both

motor types, possible interactions with FUS might be masked in a gliding assay due to lacking

accessibility of motors. Hence, both scenarios should additionally be investigated in stepping

assays, where motors are more accessible.

Axonal transport in this study was reconstituted by modifying a kinesin-1-dependent

microtubule gliding assay to be compatible with protein rich solutions such as whole cell lysates.

Cell lysates of iPSC-derived spinal motor neurons expressing wildtype FUS-GFP or

FUS-P525L-GFP did not differentially interfere with kinesin-1-dependent microtubule gliding.

However, kinesin-1 motility was more strongly impaired in the presence of cell lysates

containing endogenous 0N3R tau and supplemented with 2N4R tau-GFP compared to the

presence of 2N4R tau-GFP alone, indicating that different tau isoform levels in cell lysates can

potentially be detected using the modified kinesin-1-dependent microtubule gliding assay.

However, further evidence would be needed to confirm this hypothesis. For instance, lysates of

primary cells could be tested in this assay, e.g. using a mouse model of tauopathies (such as

FTD or progressive supranuclear palsy, where differences in tau isoform composition are a

major hallmark of the disease (Hu et al., 2007).

The precise impact of increasing 2N4R:2N3R tau isoform ratios on individual motors as

well as the contribution of the N-terminal projection domain of tau on kinesin-1 motility has to be

investigated in further stepping assays using different ratios of shorter tau isoforms (i.e. 0N, 1N).

Of particular interest in future experiments should be the interplay of these different isoforms

present at different ratios with microtubules in a gliding or stepping assay, rather than the

influence of each isoform alone.



4.6 Conclusion

Defects in axonal transport are one of the major pathological events observed early in FUS-ALS

and other neurodegenerative diseases. The exact cause and mechanism of these defects in

neurons expressing the mutant FUS-P525L variant is largely unknown to date. This study

provides evidence that neither wildtype FUS nor the mutant FUS-P525L variant directly

interferes with kinesin-1-dependent axonal transport. Further, the presented results suggest that

neither FUS variant impedes kinesin-1 motility by interacting with adaptor proteins present in

whole cell lysates of ALS patient-specific iPSC-derived spinal motor neurons.

We hence hypothesized that mislocalization of FUS-P525L to the cytoplasm corresponds

to a loss of FUS-dependent splicing activity of tau mRNA. This leads to an increased ratio of

4R:3R tau isoforms within neurons expressing this FUS mutant, affecting the kinesin-1 motility

on microtubules. In line with recent literature, the presented results show that an increased

4R:3R tau isoform ratio is indeed sufficient to impede microtubule-dependent kinesin-1 motility.

Due to the 20-fold higher microtubule binding affinity of 2N4R tau compared to 2N3R tau, the

4R tau isoform efficiently displaces the 3R tau isoform from the microtubule lattice at saturating

tau concentrations. Since 4R tau affects kinesin-1 motility more strongly than 3R tau, an

increase in the 4R:3R tau isoform ratio results in a 4R dose dependent reduction of the

observed gliding velocity.

Finally, defects in axonal transport are just one of multiple pathological processes

occurring in neurons expressing ALS-associated FUS-P525L or other mutant FUS variants.

These are not mutually exclusive, but rather interdependent and strongly connected. Hence,

one malfunctioning cellular process might affect other essential processes, thereby processively

worsening ALS pathogenesis. Whether defects in axonal transport are one of the first events

caused by mutant FUS variant expression, leading to harmful disturbances in other cellular

pathways, or whether it arises as a consequence of prior pathogenic developments, remains to

be investigated in future studies. In summary, we conclude that the axonal transport defects

seen in FUS-associated ALS arise indirectly as a consequence of FUS-P525L involvement in

one or several of the above discussed cellular mechanisms such as DNA damage repair, RNA

metabolism (especially splicing of tau), endosomal trafficking, mitochondrial homeostasis, and

stress granule dynamics.

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Mechanisms of Axonal Transport in ALS Supplementary Figures


Supplementary Figures

Figure S1: SDS-PAGE analysis of tau protein purification from insect cells.

Samples were taken at various points throughout the purification process to monitor the purification efficiency final product. SN = supernatant, after cell lysates after centrifugation and filtration; FT = flow through from HisTrap column after lysate was passed through; F1 = fraction eluted from column (first purification); AC = after cleavage, after incubation with 3C 6xHis protease; Beads = Ni-NTA beads containing the cleaved off His-tag and remaining proteins after gravity filtration; FTConc = flow through of gravity filtration containing the protein of interest and after concentrating by centrifugation; SEC = protein of interest after size exclusion chromatography

Figure S2: Different batches of insect cell expressed 4R tau cause a distinct drop in kinesin-1-dependent microtubule gliding speed at varying concentrations.

A) Kinesin-1-dependent microtubule gliding velocities in the presence of different batches of 2N4R tau with varying labels at indicated concentrations in standard motility buffer. Relative speed was calculated by dividing the median microtubule gliding velocities after 2N4R tau variant addition by the median gliding velocity of microtubules in the same channel in standard motility buffer before 2N4R tau variant addition, respectively. Averages of three independent experiments ± standard deviation are shown for the first batch of 2N4R tau-GFP and 2N4R tau-mScarlet, while relative median gliding speed ± standard deviation of one experiment are shown for 2N4R tau-mCherry and the second batch of 2N4R tau-GFP.

Mechanisms of Axonal Transport in ALS Supplementary Tables


Supplementary Tables Table S 1: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of wildtype FUS-GFP or FUS-P525L-GFP over time, BSA or standard motility buffer, corresponding to Figure 11 and Figure 12

Two-way ANOVA results: Interaction p-values: 0h 0.3194

1h 0.0983

2h 0.1869

3h 0.4816

5 nM FUS-WT-GFP FUS-P525L-GFP BSA

Buffer 0.9991 0.9885 0.9502

0h FUS-WT-GFP - 0.9977 0.9079

FUS-P525L-GFP - - 0.8285

Buffer >0.9999 >0.9999 >0.9999 1h FUS-WT-GFP - >0.9999 >0.9999

FUS-P525L-GFP - - >0.9999

Buffer >0.9999 0.9985 0.9976 2h FUS-WT-GFP - 0.9131 0.9024

FUS-P525L-GFP - - >0.9999

Buffer >0.9999 0.9975 0.9933 3h FUS-WT-GFP - 0.9968 0.9928

FUS-P525L-GFP - - >0.9999


Buffer 0.3289 0.0135 >0.9999

0h FUS-WT-GFP - 0.4325 0.2927

FUS-P525L-GFP - - 0.0111

Buffer 0.1702 0.0543 >0.9999

1h FUS-WT-GFP - >0.9999 >0.9999

FUS-P525L-GFP - - 0.5244

Buffer 0.2112 0.1338 0.8306

2h FUS-WT-GFP - 0.9952 0.7023

FUS-P525L-GFP - - 0.5587

Buffer 0.1695 0.3686 0.871

3h FUS-WT-GFP - 0.9684 0.469

FUS-P525L-GFP - - 0.7612


Buffer 0.1765 0.5127 0.9899

0h FUS-WT-GFP - 0.8327 0.0971

FUS-P525L-GFP - - 0.3302

Buffer 0.0162 >0.9999 >0.9999

1h FUS-WT-GFP - 0.3011 0.016

FUS-P525L-GFP - - 0.4378

Buffer 0.0465 0.3882 0.9997

2h FUS-WT-GFP - 0.6555 0.0375

FUS-P525L-GFP - - 0.3377

Buffer 0.0449 0.3301 0.9964

3h FUS-WT-GFP - 0.6869 0.0691

FUS-P525L-GFP - - 0.4408



Table S 2: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of 2N4R tau-GFP, corresponding to Figure 10

Two-way ANOVA results: Interaction p-value: >0.0001

1 nM 5 nM 10 nM 15 nM 25 nM 50 nM 75 nM 100 nM

0 nM >0.9999 >0.9999 >0.9999 0.8786 0.016 0.0005 0.0004 0.0004 1 nM - >0.9999 >0.9999 0.909 0.0189 0.0005 0.0005 0.0005 5 nM - - >0.9999 0.9537 0.0258 0.0007 0.0007 0.0007

10 nM - - - 0.9828 0.0366 0.001 0.001 0.001 15 nM - - - 0.9828 0.2066 0.0067 0.0063 0.0062 25 nM - - - - - 0.6482 0.6255 0.6216 50 nM - - - - - - >0.9999 >0.9999 75 nM - - - - - - - >0.9999

Table S 3: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of whole cell lysates from spinal motor neurons expressing wildtype FUS-GFP or FUS-P525L-GFP, BSA or standard motility buffer, corresponding to Figure 13B

Two-way ANOVA results: Interaction p-value: 0.0696

FUS-WT-GFP FUS-P525L-GFP BSA

Buffer 0.0691 0.0544 0.3116

50 ng/µL FUS-WT-GFP - 0.9992 0.8558

FUS-P525L-GFP - - 0.7956 Buffer 0.9973 0.3495 0.48

80 ng/µL FUS-WT-GFP - 0.2831 0.4451 FUS-P525L-GFP - - 0.998

Buffer 0.827 0.5112 0.1648 110 ng/µL FUS-WT-GFP - 0.9117 0.4083

FUS-P525L-GFP - - 0.7726 Buffer 0.9889 0.5873 0.5245

140 ng/µL FUS-WT-GFP - 0.6499 0.5884

FUS-P525L-GFP - - 0.9891

Table S 4: Significance values (p-values) for kinesin-1-depenent microtubule gliding in the presence of whole cell lysates from spinal motor neurons expressing wildtype FUS-GFP or FUS-P525L-GFP, BSA or standard motility buffer supplemented with 2N4R tau-GFP at indicated concentrations, corresponding to Figure 13C

Two-way ANOVA results: Interaction p-value: 0.6536

Tau-GFP FUS-WT-GFP FUS-P525L-GFP BSA

Buffer >0.9999 0.9876 0.9974 >0.9999

0 nM Tau-GFP - 0.9544 0.9895 >0.9999

FUS-WT-GFP - - 0.9993 0.9279

FUS-P525L-GFP - - - 0.9786

Buffer >0.9999 0.967 0.9748 >0.9999

1 nM Tau-GFP - 0.9223 0.9416 0.9991 FUS-WT-GFP - - >0.9999 0.8187

FUS-P525L-GFP - - - 0.8505



Tau-GFP FUS-WT-GFP FUS-P525L-GFP BSA

Buffer 0.9997 0.8977 0.9003 >0.9999

5 nM Tau-GFP - 0.858 0.8625 0.9989

FUS-WT-GFP - - >0.9999 0.719

FUS-P525L-GFP - - - 0.725

Buffer 0.997 0.2203 0.3247 0.9987

10 nM Tau-GFP - 0.0966 0.1882 >0.9999

FUS-WT-GFP - - 0.9978 0.0775

FUS-P525L-GFP - - - 0.1555

Buffer 0.7365 0.1383 0.0462 0.977

15 nM Tau-GFP - 0.4992 0.1674 0.8932

FUS-WT-GFP - - 0.9636 0.0992

FUS-P525L-GFP - - - 0.0182

Buffer 0.0002 <0.0001 0.0029 0.0002

25 nM Tau-GFP - 0.9991 0.7958 >0.9999

FUS-WT-GFP - - 0.6478 0.9995

FUS-P525L-GFP - - - 0.776

Buffer <0.0001 <0.0001 <0.0001 <0.0001

50 nM Tau-GFP - >0.9999 >0.9999 >0.9999

FUS-WT-GFP - - >0.9999 >0.9999 FUS-P525L-GFP - - - >0.9999

Buffer <0.0001 <0.0001 <0.0001 <0.0001

75 nM Tau-GFP - >0.9999 >0.9999 >0.9999

FUS-WT-GFP - - >0.9999 >0.9999

FUS-P525L-GFP - - - >0.9999

Buffer <0.0001 <0.0001 <0.0001 <0.0001

100 nM Tau-GFP - >0.9999 >0.9999 >0.9999

FUS-WT-GFP - - >0.9999 >0.9999

FUS-P525L-GFP - - - >0.9999

Table S 5: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of 2N3R tau-GFP and 2N4R tau-mScarlet, corresponding to Figure 17

Two-way ANOVA results: Interaction p-value: 0.9739 Concentration p-value: <0,0001 Tau variant p-value: 0.0009

3R-GFP 750 nM 1500 nM 2250 nM 3000 nM 3750 nM 4500 nM 5250 nM

500 nM 0.9999 0.7801 0.3063 0.1222 0.0082 0.0004 0.0002 750 nM - 0.9491 0.5552 0.2729 0.0245 0.0014 0.0007

1500 nM - - 0.9922 0.8952 0.265 0.0268 0.0139 2250 nM - - - 0.9996 0.7307 0.158 0.0922 3000 nM - - - 0.9449 0.3739 0.2446 3750 nM - - - - - 0.9571 0.8772 4500 nM - - - - - - >0,9999 5250 nM - - - - - - -

4R-mScarlet 150 nM 250 nM 400 nM 600 nM 800 nM 1000 nM 1200 nM

50 nM 0.6572 0.0675 0.044 0.0212 0.0008 0.0001 <0,0001

150 nM - 0.8682 0.7776 0.5955 0.0718 0.0128 0.0042 250 nM - - >0,9999 0.9997 0.6744 0.2605 0.1172



150 nM 250 nM 400 nM 600 nM 800 nM 1000 nM 1200 nM

400 nM - - - >0,9999 0.783 0.3521 0.1698 600 nM - - -

0.9151 0.5308 0.2911

800 nM - - - - - 0.996 0.9446 1000 nM - - - - - - 0.9999 1200 nM - - - - - - -

Table S 6: Significance values (p-values) for binding of 2N3R tau-GFP and 2N4R tau-mScarlet to surface-immobilized microtubules, corresponding to Figure 18

Two-way ANOVA results: Interaction p-value: 0.0001 Concentration p-value: <0,0001 Tau variant p-value: <0,0001

3R-GFP 50 nM 100 nM 200 nM 400 nM 800 nM

25 nM 0.9391 0.0013 <0,0001 <0,0001 <0,0001 50 nM - 0.0118 <0,0001 <0,0001 <0,0001

100 nM - - 0.0055 <0,0001 <0,0001 200 nM - - - 0.0975 0.0635 400 nM - - - >0,9999 800 nM - - - - -

4R-mScarlet 10 nM 250 nM 400 nM 600 nM 800 nM

5 nM 0.0067 <0,0001 <0,0001 <0,0001 <0,0001 10 nM - 0.1709 0.0206 0.0098 0.0105 25 nM - - 0.9088 0.7645 0.7807 50 nM - - - 0.9995 0.9997

100 nM - - - >0,9999 200 nM - - - - -

Table S 7: Significance values (p-values) for kinesin-1-depenent microtubule gliding in the presence of 2N3R tau-GFP and 2N4R tau-mScarlet mixed at indicated ratios, corresponding to Figure 19

One-way ANOVA results: Interaction p-value: 0.0087

4R:3R Ratio 0.33 1 3 4

0 0.2517 0.0906 0.0175 0.0076 0.33 - 0.9523 0.4437 0.2163

1 - - 0.8211 0.5184 3 - - - 0.9786 4 - - - -



Table S 8: Significance values (p-values) for binding of 2N3R tau-GFP and 2N4R tau-mScarlet mixed at indicated ratios to surface-immobilized microtubules, corresponding to Figure 20

Two-way ANOVA results: Interaction p-value: <0,0001 Ratio p-value: 0.3174 Tau variant p-value: <0,0001

3R- GFP

4R:3R Ratio 0.33 1 3 4

0 <0,0001 <0,0001 <0,0001 <0,0001 0.33 - 0.0521 0.0035 0.0008

1 - - 0.7294 0.3397 3 - - - 0.9585 4 - - - -

4R-mScarlet

4R:3R Ratio 0.33 1 3 4

0 <0,0001 <0,0001 <0,0001 <0,0001 0.33 - 0.0433 0.0008 <0,0001

1 - - 0.4112 0.0062 3 - - - 0.2277 4 - - - -

Mechanisms of Axonal Transport in ALS List of Figures


List of Figures Figure 1: Functional domains of FUS. ........................................................................................ 4 Figure 2: The motor protein kinesin-1 transports cargo along microtubules. ..............................11 Figure 3: Functional domains of tau. .........................................................................................16 Figure 4: Principle of a microtubule gliding motility assay. .........................................................20 Figure 5: Timeline for the differentiation and maturation of NPCs towards spinal motor neurons .................................................................................................................................................33 Figure 6: Assembly, mounting, and imaging of flow channels. ..................................................40 Figure 7: Reconstituting axonal transport in vitro. ......................................................................46 Figure 8: Optimal buffer conditions for reproducible kinesin-1-dependent microtubule gliding in the presence of whole cell lysates consist of 10 % glycerol, 10 nM β-glycerophosphate and 0.3 % methylcellulose in PBS. ...................................................................................................50 Figure 9: Imaging and analysis of kinesin-1-dependent microtubule gliding velocity . .............52 Figure 10: Tau-GFP binds to microtubules and inhibits kinesin-1-dependent microtubule gliding in a concentration dependent manner. ......................................................................................54 Figure 11: Recombinant FUS-GFP variants to not inhibit kinesin-1-dependent microtubule gliding. ......................................................................................................................................55 Figure 12: FUS-GFP variants do not inhibit kinesin-1-dependent microtubule gliding velocity and do not aggregate over the course of three hours. ......................................................................57 Figure 13: Cell lysates of spinal motor neurons expressing ALS-associated or wildtype FUS variants do not interfere with microtubule gliding on kinesin-1 motors. ......................................59 Figure 14: Western blot of FUS-GFP variants expressed in whole cell lysates of spinal motor neurons. ....................................................................................................................................60 Figure 15: 2N4R Tau-GFP binding to microtubules causes an arrest of microtubule gliding......62 Figure 16: 2N4R Tau-GFP binding to microtubules at low concentrations leads to an arrest in gliding of individual microtubules, while others are still motile. ..................................................63 Figure 17: The 2N4R tau isoform interferes with microtubule gliding on kinesin-1 motors at much lower concentrations compared to the 2N3R tau isoform. .........................................................65 Figure 18: The 2N4R tau-mScarlet isoform has a stronger binding affinity towards microtubules compared to 2N3R tau-GFP. .....................................................................................................67 Figure 19: An increase in 4R:3R tau isoform ratio leads to impaired kinesin-1-dependent microtubule gliding. ...................................................................................................................69 Figure 20: The 2N4R tau-mScarlet isoform prevents binding of 2N3R tau-GFP to microtubules due to its stronger binding affinity. .............................................................................................70 Figure 21: Western blot of tau variants expressed in whole cell lysates of spinal motor neurons. .................................................................................................................................................72

Mechanisms of Axonal Transport in ALS List of Tables


List of Tables Table 1: Technical equipment ...................................................................................................24 Table 2: General equipment ......................................................................................................25 Table 3: Cell culture ingredients, chemicals and reagents .........................................................26 Table 4: Media and buffers, all amounts given in percent refer to volume/volume .....................29 Table 5: Primary antibodies.......................................................................................................30 Table 6: Secondary antibodies ..................................................................................................30 Table 7: Software ......................................................................................................................31 Table 8: Characteristics for isogenic cell lines used in this study ...............................................31 Table 9: Comparison of different protein isolation techniques for their cell lysis efficiency, determined by the detection of β-actin via western blot, and protein yield, determined by BCA or Bradford assay. .........................................................................................................................48 Table 10: Calculation scheme for 4R:3R tau isoform ratios used in this study. ..........................68

Mechanisms of Axonal Transport in ALS List of Supplementary Figures and Tables


List of Supplementary Figures Figure S1: SDS-PAGE analysis of tau protein purification from insect cells. ........................... 133 Figure S2: Different batches of insect cell expressed 4R tau cause a distinct drop in kinesin-1-dependent microtubule gliding speed at varying concentrations. ............................. 133

List of Supplementary Tables Table S 1: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of wildtype FUS-GFP or FUS-P525L-GFP over time, BSA or standard motility buffer, corresponding to Figure 11 and Figure 12 ............................................................................... 134 Table S 2: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of 2N4R tau-GFP, corresponding to Figure 10 ......................................................... 135 Table S 3: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of whole cell lysates from spinal motor neurons expressing wildtype FUS-GFP or FUS-P525L-GFP, BSA or standard motility buffer, corresponding to Figure 13B .................... 135 Table S 4: Significance values (p-values) for kinesin-1-depenent microtubule gliding in the presence of whole cell lysates from spinal motor neurons expressing wildtype FUS-GFP or FUS-P525L-GFP, BSA or standard motility buffer supplemented with 2N4R tau-GFP at indicated concentrations, corresponding to Figure 13C ........................................................... 135 Table S 5: Significance values (p-values) for kinesin-1-dependent microtubule gliding in the presence of 2N3R tau-GFP and 2N4R tau-mScarlet, corresponding to Figure 17 ................... 136 Table S 6: Significance values (p-values) for binding of 2N3R tau-GFP and 2N4R tau-mScarlet to surface-immobilized microtubules, corresponding to Figure 18 ........................................... 137 Table S 7: Significance values (p-values) for kinesin-1-depenent microtubule gliding in the presence of 2N3R tau-GFP and 2N4R tau-mScarlet mixed at indicated ratios, corresponding to Figure 19 ................................................................................................................................. 137 Table S 8: Significance values (p-values) for binding of 2N3R tau-GFP and 2N4R tau-mScarlet mixed at indicated ratios to surface-immobilized microtubules, corresponding to Figure 20 .... 138

Mechanisms of Axonal Transport in ALS Summary


Summary Background

Neurodegenerative diseases have become one of the most common causes of death worldwide

over the last couple of decades, with increasing tendency. Amyotrophic lateral sclerosis (ALS) is

the most common neurodegenerative disease affecting specifically spinal (lower) and cortical

(upper) motor neurons in the spinal cord and brainstem, respectively. It is usually a late onset

disorder (average age of onset in Germany is 61 years) and leads to death within 2-5 years

after symptoms onset due to respiratory failure. To date, there is no cure for ALS and only two

drugs have been approved for its treatment, which prolong the lifespan for up to six months or

slow down disease progression in a subpopulation of patients. About 90 % of ALS cases are

sporadic, while about 10 % are familial and hence caused by mutations in specific genes,

among them fused in sarcoma (FUS), a DNA- and RNA-binding protein. Mutations in FUS

account for roughly 5 % of familial cases and occur predominantly in its nuclear localization

sequence (NLS), such as the FUS-P525L mutation. Neurons expressing this variant display a

strong cytoplasmic mislocalization of FUS and hence a loss of its nuclear function. Among other

pathological events, defects in axonal transport along microtubules have been observed early in

disease progression in several models of FUS-ALS, indicating its role as a major hallmark of the

disease. However, the mechanism of how transport is impaired within these neurons remains

unknown to date.

Hypothesis

This study aimed at investigating two possible mechanisms how the FUS-P525L mutant variant

affects microtubule-based axonal transport. First, it was analyzed whether FUS directly interacts

with microtubules or motors and if the mislocalized, mutant variant alters this interaction.

Secondly, cytoplasmic mislocalized FUS-P525L can no longer fulfil its regular role in the splicing

of pre-mRNAs, among them the mRNA coding for the microtubule-associated protein tau. This

reportedly leads to an increased ratio of translated tau isoforms containing four microtubule

binding repeats (4R) to those containing three repeats (3R). 4R tau isoforms are known to have

a stronger binding affinity towards microtubules and may hence impair transport more severely

by acting as a roadblock for motor proteins. Towards this end, this study investigated whether

an increase in 4R:3R tau isoform ratio is sufficient to impair microtubule based transport.

Methods

Axonal transport was reconstituted in vitro using a kinesin-1-dependent microtubule gliding

assays, in which microtubules are propelled by surface-immobilized kinesin-1 motors. The

assay was modified and optimized to operate sensitively and robust in the presence of complex

solutions such as whole cell lysates and the microtubule gliding velocity analyzed as a measure

for motility of the underlying motors. To determine the direct interaction of FUS variants with

Mechanisms of Axonal Transport in ALS Summary


kinesin-1 or microtubules, recombinant human wildtype FUS-GFP and FUS-P525L-GFP was

added to the assay. In addition, ALS patient-specific induced pluripotent stem cells (iPSCs)

expressing the same FUS variants were differentiated towards spinal motor neurons and their

cell lysates applied to this assay in order to determine whether FUS variants need endogenous

adaptors or interaction partners to interfere with kinesin-1 motility on microtubules. Further, to

investigate the interference of tau isoforms with kinesin-1 motility, recombinant human 2N3R

tau-GFP and 2N4R tau-mScarlet was purified from insect cells and added to the modified

kinesin-1-dependent microtubule gliding assay, either individually or combined at different

ratios. In addition, the binding of these tau variants to microtubules was assessed.

Results

The kinesin-1-dependent microtubule gliding assays was modified to operate sensitively and

robustly in the presence of β-glycerophosphate (to inhibit endogenous phosphatases in whole

cell lysates), and methylcellulose (to prevent microtubule detachment from kinesin-1 motors due

to presence of β-glycerophosphate). Under these conditions, neither recombinant human

FUS-GFP nor endogenous FUS-GFP variants in lysates of spinal motor neurons bound to

microtubules or interfered with kinesin-1 motility. In contrast, both tau isoforms used in the

present study bound to microtubules and impaired kinesin-1 motility, while 2N4R tau-mScarlet

was a much more potent inhibitor of microtubule gliding and displayed a 20-fold stronger binding

affinity to microtubules compared to 2N3R tau-GFP. Interestingly, increasing ratios of 4R:3R tau

isoforms impaired kinesin-1-dependent microtubule gliding. In addition, the presence of 2N4R

tau-mScarlet strongly prevented 2N3R tau-GFP from binding to microtubules.

Conclusion

This study provides evidence that neither wildtype FUS nor the FUS-P525L variant directly

interfere with axonal transport by interacting with kinesin-1 motors or microtubules. Further, the

present data suggests that neither FUS variant impedes kinesin-1 motility on microtubules by

interacting with endogenous adaptor proteins present in cell lysates of iPSC-derived spinal

motor neurons. Therefore, it is proposed that axonal transport defects are not directly caused by

interaction of cytoplasmic mislocalized FUS with the motors or microtubules, but rather arise as

a consequence of other pathological events triggered by mutant FUS variants. In particular, this

study demonstrates that an increased ratio of 4R:3R tau isoforms is sufficient to impair kinesin-1

motility on microtubules due to increased decoration of microtubules with 4R tau isoforms,

preventing 3R tau isoforms from binding to microtubules. This strongly suggests that an

increased ratio of 4R:3R tau isoforms, since FUS no longer regulates splicing of tau pre-mRNA

upon its cytoplasmic mislocalization, may be sufficient to cause or contribute to the axonal

transport defects observed early in FUS-ALS pathology.

Mechanisms of Axonal Transport in ALS Zusammenfassung


Zusammenfassung Hintergrund

Neurodegenerative Erkrankungen sind in den letzten Jahrzehnten mit zunehmender Tendenz

zu einer der häufigsten Todesursachen weltweit geworden. Amyotrophe Lateralsklerose (ALS)

ist die häufigste neurodegenerative Erkrankung, die spezifisch spinale (untere) und kortikale

(obere) Motoneuronen im Rückenmark bzw. im Hirnstamm betrifft. Es handelt sich in der Regel

um eine spät einsetzende Krankheit (das mittlere Erkrankungsalter in Deutschland beträgt 61

Jahre) und führt innerhalb von 2-5 Jahren nach Auftreten der Symptome zum Tod aufgrund von

Atemversagen. Bisher gibt es keine Heilung für ALS und es wurden nur zwei Medikamente für

die Behandlung zugelassen, die die Lebensdauer um bis zu sechs Monate verlängern oder das

Fortschreiten der Krankheit bei einer Subpopulation von Patienten verlangsamen. Ungefähr

90% der ALS-Fälle sind sporadisch, während ungefähr 10% familiär sind und daher durch

Mutationen in bestimmten Genen verursacht werden, darunter fused in sarcoma (FUS), einem

DNA- und RNA-bindenden Protein. Mutationen in FUS machen etwa 5% der familiären Fälle

aus und treten überwiegend in der Kernlokalisierungssequenz (NLS) auf, wie beispielsweise die

FUS-P525L Mutation. Neuronen, die diese Mutante exprimieren, zeigen eine starke

zytoplasmatische Fehllokalisierung von FUS und damit einen Verlust seiner Funktionen im

Zellkern. Neben anderen pathologischen Ereignissen wurden in mehreren FUS-ALS

Modellsystemen Defekte im Mikrotubuli-basierenden axonalen Transport früh im

Krankheitsverlauf beobachtet, was auf seine Rolle als eines der Hauptmerkmale dieser

Krankheit hindeutet. Der Mechanismus, wie der Transport innerhalb dieser Neuronen

beeinträchtigt wird, ist jedoch bis heute unbekannt.

Hypothese

Ziel dieser Studie ist es, zwei mögliche Mechanismen zu untersuchen, wie das mutierte

FUS-P525L Protein den axonalen Transport entlang von Mikrotubuli beeinflusst. Zunächst

wurde analysiert, ob FUS direkt mit Mikrotubuli oder Motorproteinen interagiert und ob

zytoplasmatische fehllokalisierte FUS-P525L Protein diese Interaktion verändert. Ferner kann

zytoplasmatische fehllokalisiertes FUS-P525L seine reguläre Rolle beim Spleißen von

Prä-mRNAs nicht mehr erfüllen, darunter die mRNA, die für das mit Mikrotubuli-assoziierte

Protein Tau kodiert. Dies führt zu einem erhöhten Verhältnis von translatierten Tau-Isoformen,

die vier Mikrotubuli-Bindestellen (4R) enthalten, zu solchen mit drei Bindestellen (3R). Es ist

bekannt, dass 4R-Tau-Isoformen eine stärkere Bindungsaffinität zu Mikrotubuli im Vergleich zu

3R-Tau-Isoformen aufweisen und daher den Transport stärker beeinträchtigen können, indem

sie als Hindernis für Motorproteine agieren. In dieser Studie wurde daher untersucht, ob eine

Erhöhung des Verhältnisses von 4R:3R-Tau-Isoform ausreicht, um den Mikrotubuli-basierenden

Transport zu beeinträchtigen.



Methoden

Der axonale Transport wurde in vitro unter Verwendung eines Kinesin-1-gestuerten Mikrotubuli

Motilitätsassay rekonstruiert, bei welchem Mikrotubuli von darunterliegenden

oberflächenimmobilisierte Kinesin-1 Motorproteinen transportiert werden, also über die

Oberfläche gleiten. Der Assay wurde modifiziert und optimiert, um in Gegenwart komplexer

Lösungen wie Ganzzelllysaten sensitiv und robust zu funktionieren, und die

Gleitgeschwindigkeit der Mikrotubuli wurde als Maß für die Motilität der darunterliegenden

Motoren analysiert. Um die direkte Wechselwirkung von FUS-Varianten mit Kinesin-1

Motorproteinen oder Mikrotubuli zu bestimmen, wurde dem Assay rekombinantes menschliches

Wildtyp-FUS-GFP und FUS-P525L-GFP hinzugegeben. Zusätzlich wurden ALS-

patientenspezifische, induzierte pluripotente Stammzellen (iPSCs), welche dieselben FUS-

Varianten exprimieren, zu spinalen Motoneuronen differenziert und ihre Zelllysate in diesem

Assay angewendet, um zu bestimmen, ob FUS-Varianten endogene Adapter oder

Interaktionspartner für die Interaction mit Kinesin-1 oder Mikrotubuli benötigen. Um den Einfluss

von Tau-Isoformen auf die Kinesin-1 Motilität zu untersuchen, wurde rekombinantes

menschliches 2N3R Tau-GFP und 2N4R Tau-mScarlet aus Insektenzellen aufgereinigt und

dem modifizierten Kinesin-1-gestuerten Mikrotubuli Motilitätsassay entweder einzeln oder in

unterschiedlichen Verhältnissen kombiniert hinzugegeben. Zusätzlich wurde die Bindung dieser

Tau-Varianten an Mikrotubuli analysiert.

Ergebnisse

Die Kinesin-1-gesteuerte Mikrotubuli Motilitätsassay wurden so modifiziert, dass er in

Gegenwart von β-Glycerophosphat (zur Hemmung endogener Phosphatasen in

Ganzzelllysaten) und Methylcellulose (zur Verhinderung der Ablösung von Mikrotubuli von

Kinesin-1 Motoren aufgrund von β-Glycerophosphat) empfindlich und robust funktioniert. Unter

diesen Bedingungen zeigten weder rekombinantes menschliches FUS-GFP noch endogene

FUS-GFP-Varianten in Lysaten von spinalen Motoneuronen eine Wechselwirkung mit

Mikrotubuli und beeinträchtigten auch nicht die Kinesin-1 Motilität. Im Gegensatz dazu banden

beide in der vorliegenden Studie verwendeten Tau-Isoformen an Mikrotubuli und

beeinträchtigten die Kinesin-1-Motilität, wobei 2N4R Tau-mScarlet das Gleitens von Mikrotubuli

viel stärkerer beeinträchtigte und eine 20-fach stärkere Bindungsaffinität zu Mikrotubuli im

Vergleich zu 2N3R Tau-GFP zeigte. Ferner beeinträchtigten steigende Verhältnisse von 4R:3R

Tau-Isoformen über Kinesin-1 gleitende Mikrotubuli, während die Präsenz von 2N4R

Tau-mScarlet die Bindung von 2N3R Tau-GFP an Mikrotubuli stark verminderte.

Schlussfolgerung

Diese Studie liefert Hinweise darauf, dass weder Wildtyp-FUS noch die FUS P525L-Variante

den axonalen Transport direkt beeinflussen, da sie nicht mit Kinesin-1 Motorproteinen oder

Mikrotubuli interagieren. Ferner legen die vorliegenden Daten nahe, dass keine der FUS-



Varianten die Kinesin-1 Motilität auf Mikrotubuli durch Wechselwirkung mit endogenen

Adapterproteinen behindert, die in Zelllysaten von iPSC-differenzierte spinalen Motoneuronen

vorhanden sind. Dies legt nahe, dass axonale Transportdefekte nicht durch direkte

Wechselwirkung von zytoplasmatisch fehllokalisiertem FUS Protein mit Motorproteinen oder

Mikrotubuli verursacht werden, sondern als Folge anderer pathologischer Ereignisse auftreten,

die durch mutierte FUS-Varianten entstehen. Insbesondere zeigt diese Studie, dass ein

erhöhtes Verhältnis von 4R:3R Tau-Isoformen ausreicht, um die Kinesin-1 Motilität auf

Mikrotubuli zu behindern. Dies geschieht vermutlich aufgrund der erhöhten Bindung von 4R

Tau-Isoformen an Mikrotubuli, weil dadurch die Bindung von 3R Tau-Isoformen an Mikrotubuli

verhindert wird. Dies deutet stark darauf hin, dass ein erhöhtes Verhältnis von 4R:3R

Tau-Isoformen, verursacht durch die fehlende regulatorische Beteiligung von FUS am Spleißen

von Tau-Prä-mRNA aufgrund der zytoplasmatischen Fehllokalisation von FUS, wahrscheinlich

zu den axonalen Transportdefekten beiträgt, die früh in der FUS-ALS-Pathologie beobachtet

wurden.

Mechanisms of Axonal Transport in ALS Acknowledgements


Acknowledgements I would like to thank Prof. Dr. Dr. Andreas Hermann and Prof. Dr. Stefan Diez for the

opportunity to pursue my doctoral studies in their labs. I wish to thank them for their continuous

support, constructive criticism, scientific input and motivation during the last years. Further, I

would like to thank Prof. Kempermann, Prof. Falkenburger, and Prof. Karl for their participation

in my thesis committee.

This work was supported in part by the NOMIS foundation, German Federal Ministry of

Education and Research BMBF OptiZeD 03Z22E511, by a seed grant of the Center For

Regenerative Therapies Dresden (CRTD), the Helmholtz Virtual Institute “RNA dysmetabolism

in ALS and FTD (VH-VI-510)”, an unrestricted grant by a family of a deceased ALS patient, and

the Stiftung zur Förderung der Hochschulmedizin in Dresden, and I am very grateful for the

support of all these funding sources.

Additionally, I would like to thank Dr. Jie Wang from the Max Planck Institute of Molecular

Cell Biology and Genetics, Dresden, for providing recombinant human FUS-GFP variant protein,

and Dr. Marcus Braun from the Institute of Biotechnology CAS, Prague, for the first batch of

recombinant human 2N4R tau-GFP protein. In line with this, I would like to thank Prof. Eckhardt

Mandelkow at the German Center for Neurodegenerative Diseases e.V. Bonn for the provision

of a DNA construct coding for recombinant human 2N3R tau-GFP and Dr. Amayra Hernandez

Vega for the construct coding for 2N4R tau-GFP. I am extremely grateful to Dr. Jens Ehrig from

the Molecular Imaging and Manipulation facility at B CUBE, who always took the time to

optimize my imaging workflows in the best way possible.

Further, I would like to voice my heartfelt appreciation to everyone who supported me in

one way or another during the course of my thesis. My deepest gratitude goes to Dr. Hauke

Drechsler, who significantly contributed to the success of this project through his scientific input,

but also through his continuous motivation (both verbally and with chocolate) and cure of panic

attacks. On the same note, I’m very grateful to Corina Bräuer for her unconditional support in

the lab, but also for always having an open ear and looking after me during tough times.

Together with Nora Hartmann, we always had a fun time in the lab, especially (but not

exclusively) on Fridays, and I will miss our almost philosophic discussions.

A huge thank you goes to Dr. Rahul Grover for his help in generating recombinant human

2N4R tau-mScarlet and 2N3R tau-GFP constructs and for his help and supervision during the

purification process. Likewise, my sincere gratitude goes to all Diez lab members who

supported me and this project during the last four years for the fruitful discussions and fun times

Mechanisms of Axonal Transport in ALS Acknowledgements


in and outside the lab: Dr. Till Korten, Foram Joshi, Dr. Veikko Geyer, Dr. Cordula Reuther,

Ashwin D’ Souza, Laura Meißner. I would also like to point out the amazing Barbara Lindemann

and thank her for her intensive administrational support and help with every not science-related

question.

I also would like to thank former and current members of the Hermann lab for their

continuous support in the lab and our fun lab nights at various restaurants and Christmas

markets: Barbara Szewczyk, Marie Anskat, Dr. Hannes Glaß, Yu Niu, Marcel Naumann,

Dr. Arun Pal, and Dr. Anne-Karen Meyer. In particular, I would like to thank Anett, Sylvia,

Andrea, Katja and Conny for their help with all organizational matters in the lab and their

sympathy. Special appreciation goes to Dr. Julia Japtok for her scientific, but especially her

emotional support throughout the last four years, even after she finished her doctoral thesis.

Schließlich möchte ich mich bei meiner Familie für ihre unendliche Unterstützung während

meiner Doktorzeit bedanken. Besonders danke ich dabei meiner Astaoma, die mich jederzeit

bei sich willkommen geheißen hat, mit und ohne Wäsche, mit und ohne Nala.

Außerdem danke ich meinem Freund Tino dafür, dass er mir in stressigen Zeiten viel

abgenommen und mich unterstützt hat, wann immer er konnte.

Vor allem aber danke ich meiner Mama für ihre bedingungslose Unterstützung während

meiner gesamten Ausbildung. Du hattest immer ein offenes Ohr, hast immer eine Lösung

gefunden oder wusstest Rat, und du hast mir immer Mut zum Weitermachen gegeben. Du hast

mir die Kraft gegeben bis hierhin und noch viel weiter zu kommen, und dafür danke ich dir von

ganzem Herzen.

Mechanisms of Axonal Transport in ALS Anlage 1


Anlage 1 Erklärungen zur Eröffnung des Promotionsverfahrens,

Technische Universität Dresden

Medizinische Fakultät Carl Gustav Carus

Promotionsordnung vom 24. Juli 2011

1. Hiermit versichere ich, dass ich die vorliegende Arbeit ohne unzulässige Hilfe Dritter und

ohne Benutzung anderer als der angegebenen Hilfsmittel angefertigt habe; die aus fremden

Quellen direkt oder indirekt übernommenen Gedanken sind als solche kenntlich gemacht.

2. Bei der Auswahl und Auswertung des Materials sowie bei der Herstellung des Manuskripts

habe ich Unterstützungsleistungen von folgenden Personen erhalten:

Prof. Dr. Dr. Andreas Hermann

Prof. Dr. Stefan Diez

Dr. Hauke Drechsler

3. Weitere Personen waren an der geistigen Herstellung der vorliegenden Arbeit nicht beteiligt.

Insbesondere habe ich nicht die Hilfe eines kommerziellen Promotionsberaters in Anspruch

genommen. Dritte haben von mir weder unmittelbar noch mittelbar geldwerte Leistungen für

Arbeiten erhalten, die im Zusammenhang mit dem Inhalt der vorgelegten Dissertation stehen.

4. Die Arbeit wurde bisher weder im Inland noch im Ausland in gleicher oder ähnlicher Form

einer anderen Prüfungsbehörde vorgelegt.

5. Die Inhalte dieser Dissertation wurden in folgender Form veröffentlicht: Noch nicht zutreffend.

6. Ich bestätige, dass es keine zurückliegenden erfolglosen Promotionsverfahren gab.

7. Ich bestätige, dass ich die Promotionsordnung der Medizinischen Fakultät der Technischen

Universität Dresden anerkenne.

8. Ich habe die Zitierrichtlinien für Dissertationen an der Medizinischen Fakultät der

Technischen Universität Dresden zur Kenntnis genommen und befolgt.

9. Ich bin mit den "Richtlinien zur Sicherung guter wissenschaftlicher Praxis, zur Vermeidung

wissenschaftlichen Fehlverhaltens und für den Umgang mit Verstößen" der Technischen

Universität Dresden einverstanden.

Ort, Datum Unterschrift des Doktoranden

Mechanisms of Axonal Transport in ALS Anlage 2


Anlage 2 Erklärung über die Einhaltung gesetzlicher Bestimmungen.

Hiermit bestätige ich die Einhaltung der folgenden aktuellen gesetzlichen Vorgaben im Rahmen

meiner Dissertation

Untersuchungen mit Personenbezug oder Sachverhalten, die das Medizinproduktegesetz

betreffen

Aktenzeichen der zuständigen Ethikkommission: EK45022009; EK393122012

Entfällt

Az. 55-8811.71/170

mungen der Medizinischen Fakultät und des

Universitätsklinikums Carl Gustav Carus.

Ort, Datum Unterschrift des Doktoranden

Mechanisms of Axonal Transport in ALS

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