EXAMINING DYNAMIC ASPECTS OF PRESYNAPTIC TERMINAL FORMATION
VIA
LIVE CONFOCAL MICROSCOPY
By
LUKE ANDREW DASCENZO BURY
Submitted in partial fulfillment of the requirements
For the degree of Doctor of Philosophy
Dissertation Adviser: Dr. Shasta Sabo
Department of Pharmacology
CASE WESTERN RESERVE UNIVERSITY
August, 2015
II
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the dissertation of
Luke A. D. Bury
candidate for the Doctor of Philosophy degree*.
Committee Chair
Ruth Siegel
Committee Member
Shasta Sabo
Committee Member
Lynn Landmesser
Committee Member
Brian McDermott
Committee Member
Edwin Levitan
Date of Defense
June 7th
, 2015
*We also certify that written approval has been obtained
For any proprietary material contained therein
III
Table of Contents
List of Tables VII
List of Figures VIII
Abstract X
Chapter 1: General Introduction 1
Basic synaptic physiology 2
Presynaptic distribution and composition 3
STVs and PTVs – transport vesicles for efficient presynaptic protein delivery 4
STV and PTV recruitment into development synapses – a dynamic process 12
Synaptogenic signaling through trans-synaptic adhesion proteins 14
Chapter 2: Coordinated trafficking of synaptic vesicle and active zone proteins prior
to synapse formation 24
Abstract 25
Introduction 26
Results 29
STVs and PTVs can move together 29
STVs and PTVs display similar pausing characteristics 33
IV
STVs and PTVs pause simultaneously at the same sites in the axon 36
Recruitment of a PTV enhances accumulation of additional PTVs
at sites of synapse formation 44
STV pausing is only mildly dependent on PTVs 46
Materials and Methods 51
Acknowledgements 55
Chapter 3: Dynamic mechanisms of neuroligin-dependent presynaptic terminal
assembly in living cortical neurons 56
Abstract 57
Introduction 58
Results 60
Synaptic protein recruitment fluctuates frequently and rapidly during
assembly of individual presynaptic terminals 60
Levels of synaptic vesicle proteins are highly labile at developing
axo-dendritic contacts 69
Synaptophysin is preferentially stabilized at clustered neurexin 72
Actin polymerization is occasionally co-localized with neurexin
clustering 76
Levels of recruited synaptophysin and neurexin are strongly
correlated at developing presynaptic terminals 77
V
Trans-synaptic signaling recruits synaptic proteins without
substantially altering their transport 80
Synaptic vesicle and active zone protein transport vesicles are not
attracted to sites of trans-synaptic signaling 85
Initial arrival of STVs and PTVs to a developing synapse is likely
uncoordinated 91
Materials and Methods 93
Acknowledgements 98
Chapter 4: Imaging presynaptic dynamics during mouse cortical development via
two-photon confocal microscopy 99
Introduction 100
Results 101
Materials and Methods 111
Chapter 5: Discussion 115
Is STV and PTV transport coordinated? 115
What causes STV and PTV pausing? 117
Why don’t all vesicles pause at any given site? 119
A model of coordinated transport 120
VI
What are the dynamics of presynaptic protein recruitment at developing
terminals? 122
Presynaptic proteins are trapped at, but not attracted to, developing synapse 122
Synaptic proteins are labile at developing synapses 124
A model of neuroligin-neurexin mediated presynaptic protein recruitment 125
A model for dynamic presynaptic protein recruitment to a developing
synapse 127
In vivo synaptic dynamics 128
Future outlooks – directions for future investigations 130
Conclusion 134
Bibliography 135
VII
List of Tables
Table 1: STV and PTV proteins 6
Table 2: Trans-synaptic adhesion partners 17
VIII
List of Figures
Figure 1.1: Newly synthesized presynaptic proteins are packaged into STVs and PTVs
and then transported along axonal microtubules for delivery to developing
synapses 7
Figure 1.2: Mechanisms of regulation of STV and PTV transport 11
Figure 1.3: Recruitment of presynaptic cargo downstream of neurexin dependent
synaptogenic adhesion 21
Figure 2.1: STVs and PTVs move together 31
Figure 2.2: PTV pausing is qualitatively and quantitatively similar to STV pausing 34
Figure 2.3: STVs and PTVs pause at the same sites 38
Figure 2.4: STVs and PTVs preferentially pause at the same sites at the same time 42
Figure 2.5: Multiple PTVs are attracted to the same sites 46
Figure 2.6: A direct interaction between STVs and PTVs cannot account for the
attraction of STVs to pause sites that contain PTVs 49
Figure 3.1: Levels of synaptic vesicle protein enrichment at individual trans-synaptic
adhesion sites are modulated throughout recruitment 63
Figure 3.2: Modulation of synaptic vesicle protein recruitment occurs through two
distinct mechanisms 67
Figure 3.3: Synaptic vesicle protein recruitment to axo-dendritic contacts is similar
that at induced trans-synaptic signaling sites 70
IX
Figure 3.4: Synaptic vesicle protein accumulation occurs at clustered neurexin
and correlates with levels of clustered neurexin 74
Figure 3.5: Neurexin and synaptophysin levels fluctuate over the course of minutes
at axo-dendritic contact sites and neuroligin induced developing synapses 79
Figure 3.6: Trans-synaptic signaling has little effect on synaptic vesicle or active
zone protein transport 83
Figure 3.7: Trans-synaptic signaling does not attract synaptic vesicle or active zone
proteins 89
Figure 4.1: Synaptophysin-tdtomato expression in transgenic knock-in mouse 103
Figure 4.2: Synaptophysin-tdtomato is localized to presynaptic terminals 105
Figure 4.3: Synaptophysin-tdtomato puncta both split and merge in vivo 108
Figure 4.4: Instantaneous velocity and mean displacement decrease as
development progresses 110
Figure 5.1: Models for coordination of recruitment of STVs and PTVs to the same
place at the same time 121
Figure 5.2: Model of synaptic vesicle protein transport vesicle (STV) recruitment
to a developing presynaptic terminal 128
X
Examining Dynamic Aspects of Presynaptic Terminal
Formation via Live Confocal Microscopy
Abstract
By
LUKE ANDREW DASCENZO BURY
To create a presynaptic terminal, molecular signaling events must be orchestrated within
a number of subcellular compartments. In the soma, presynaptic proteins need to be
synthesized, packaged together, and attached to microtubule motors for shipment through
the axon. Within the axon, transport of presynaptic packages is regulated in order to
ensure that developing synapses receive an adequate supply of components. At individual
axonal sites, extracellular interactions must be translated into intracellular signals that can
incorporate mobile transport vesicles into the nascent presynaptic terminal. Even once the
initial recruitment process is complete, the components and subsequent functionality of
presynaptic terminals need to constantly be remodeled. Perhaps most remarkably, all of
these processes need to be coordinated in space and time. In this dissertation, I will
discuss how these dynamic cellular processes occur in neurons of the central nervous
system in order to generate presynaptic terminals in the brain, and describe experiments
to further elucidate these mechanisms.
1
Chapter 1: General Introduction
Over 100 years ago, seminal work by Ramon y Cajal led to the development of
the neuron doctrine, which states that the nervous system is comprised of individual cells
(1, 2). Inspired by Cajal, Charles Sherrington proposed the existence of synapses that
would allow communication between these cells (2, 3). From these early beginnings, it
has become clear that the synaptic connections between neurons are indispensible for
proper nervous system function and for life itself.
Synaptogenesis is important throughout life. Extensive synapse formation during
neonatal and early postnatal development is responsible for initial establishment of the
neural circuitry in the brain. However, synapse formation does not cease upon
maturation. Synaptogenesis, in concert with synapse elimination, is a constant process
that occurs throughout life as circuits in the brain are modified in response to varying
external stimuli. The ability of the brain to constantly remodel its circuitry over time is
critical in order to successfully adapt to and live within changing environments over the
course of an organism’s life.
Studying how synapses form is necessary to understand both how the brain
functions normally and how this goes awry in disease. Aberrations in synaptogenesis
have been linked to a number of neurodevelopmental disorders, including autism and
schizophrenia (4-7). In addition, poor synapse formation has been implicated in a wide
variety of non-developmental neurological diseases, including addiction and depression
(8-11). Furthermore, synapse loss is one of the hallmarks of neurodegenerative diseases
such as Alzheimer’s disease, Parkinson’s disease, and Huntington’s disease (12-15).
While it is not clear whether synapse formation is decreased or synapse elimination is
2
enhanced in these diseases (or both), enhancing synaptogenesis is an important
therapeutic strategy to mitigate the effects of these devastating diseases (16).
Basic synaptic physiology
The basic purpose of a synapse in the brain is to enable transmission of electrical
signals from one neuron to another. There are two types of synapses in the brain that
accomplish this feat: electrical and chemical. Electrical synapses enable a direct
connection between two neurons through the formation of gap junctions (17, 18). Gap
junctions are specialized connections mediated by connexin proteins that result in a pore
forming between two connected cells (19, 20). This pore not only physically links the
cells, it also electrically links them by allowing rapid diffusion of ions from the
cytoplasm in either direction across the junction. Therefore, unlike chemical synapses,
electrical information can be transmitted by either cell to its connected partner. Because
of this near-instantaneous bi-directional signaling capability, electrical synapses are often
found between inhibitory neurons in the brain where they are utilized to synchronize
inhibitory signals (17, 21).
While critical for normal nervous system function, electrical synapses make up
only a small portion of synapses in the brain. Chemical synapses take the role as the
major synaptic type in both the peripheral and central nervous systems. At chemical
synapses, cells are physically separated by the synaptic cleft and electrical information is
transferred between neurons via neurotransmitters. As action potentials reach presynaptic
terminals, the resulting depolarization opens voltage-gated calcium channels within the
presynaptic plasma membrane, leading to an influx of calcium. This, in turn, causes
synaptic vesicles (SVs) loaded with neurotransmitter to fuse with the active zone
3
membrane, via exocytic SNARE proteins. As SVs fuse, neurotransmitters are released
into the synaptic cleft, which stimulates the postsynaptic cell through the opening of
neurotransmitter-gated ion channels and/or the activation of metabotropic
neurotransmitter receptors. For more thorough reviews of presynaptic function please see
the following (22-25).
The neurotransmitter and postsynaptic neurotransmitter receptors vary according
to the neuronal cell type and location. In the brain, glutamate is the main excitatory
neurotransmitter, while gamma-aminobutyric acid (GABA) functions as the main
inhibitory neurotransmitter. However, many other neurotransmitters are produced and
released in the brain, including dopamine, serotonin, acetylcholine, norepinephrine, ATP,
cannabinoids, and various peptides (26-33). These other neurotransmitters can function in
either an excitatory or inhibitory role, depending on the circuit they are utilized in and the
type of receptor they activate.
Presynaptic distribution and composition
The location and number of presynaptic terminals within an axon varies. For
motor neurons, presynaptic structures known as terminaux boutons are located at the
distal end of the projecting axon where it makes contact with the muscle. In contrast,
locally projecting neurons of the cerebral cortex typically form “en passant” presynaptic
terminals along much of the length of unmyelinated axons. Strikingly, the number of
presynaptic terminals per axon in the cortex can number in the thousands (34-36).
Recently, an elegant study showed that an “average” presynaptic terminal in the
cortex or cerebellum is composed of approximately 300,000 individual molecules (37).
4
Although the number and diversity of proteins at the synapse is striking, many
presynaptic proteins can be classified into two important groups: (i) synaptic vesicle (SV)
associated proteins and (ii) proteins of the cytomatrix at the active zone (CAZ) (38, 39).
SVs are 35-45nm spherical vesicles that fuse with the synaptic plasma membrane to
release neurotransmitter (25, 40-43). SV-associated proteins control neurotransmitter
uptake into SVs, trafficking of SVs within the synapse, and fusion of SVs with the
plasma membrane. CAZ proteins are structural components of the presynaptic terminal,
tethering SVs to the active zone and priming them for fusion with the plasma membrane.
In addition, SNARE proteins within the CAZ interact with the v-SNARE
synaptobrevin/VAMP2 on synaptic vesicles to enable SV fusion with the plasma
membrane (44).
STVs and PTVs – transport vesicles for efficient presynaptic protein delivery
Building presynaptic terminals in the CNS poses unique challenges for the
neuron. Most of the proteins that comprise the presynaptic side of the synapse are
translated in the soma. Since some axons extend for millimeters, presynaptic components
oftentimes need to be transported long distances from where they were made.
Furthermore, presynaptic terminals form along an individual axon in an unsynchronized
manner. This requires specific assembly of presynaptic proteins and structures at the right
place at the right time.
To facilitate efficient delivery to developing synapses, groups of presynaptic
components are sorted into precursor organelles (Figure 1.1). Specifically, SV proteins
are incorporated into Synaptic vesicle precursor protein Transport Vesicles (STVs), while
CAZ proteins are packaged into Piccolo-bassoon Transport Vesicles (PTVs) (45, 46).
5
STVs are tubulo-vesicular structures that are heterogeneous in size and shape (47-49).
These golgi-derived vesicles can cycle with the axonal membrane and primarily contain
proteins that are associated with SVs, including synapsin, synaptophysin, SV2, VGlut1,
VAMP2, Rab3a, synaptotagmin and amphiphysin (Table 1) (47-58). PTVs primarily
exist as 80nm dense-core vesicles (59, 60) that are derived from the trans-golgi network
(61, 62), and they carry proteins of the CAZ, including piccolo, bassoon, syntaxin, RIM,
Munc-18, ELKS2/CAST, SNAP-25, and n-cadherin (Table 1) (49, 59-62).
6
Table 1: STV and PTV proteins
Synaptic proteins Motors Motor Linkage
STVs
Synaptic vesicle proteins, including:
synapsin, synaptophysin, SV2, VGlut1,
VAMP2, Rab3a, synaptotagmin and
amphiphysin
kinesin-3/KIF1A
and KIF1B
syd-2/liprin-
DENN/MADD, via
Rab 3
SAM-4
PI(4,5)P(2)
kinesin-1/KIF5 ?
dynein dynactin complex
PTVs
Active zone cytomatrix proteins, including:
piccolo, bassoon, syntaxin, RIM, Munc-18,
ELKS2/CAST, SNAP-25, and n-cadherin
kinesin-1/KIF5 syntabulin, via
syntaxin
Dynein bassoon
7
Figure 1.1: Newly synthesized presynaptic proteins are packaged into STVs and
PTVs and then transported along axonal microtubules for delivery to developing
synapses. STVs and PTVs are derived from the trans-Golgi network. CAZ and SV-
associated proteins are sorted into PTVs and STVs, respectively. STVs and PTVs then
attach to kinesins and potentially dynein microtubule (MT) motors. Attachment can occur
through direct binding of motors and their cargo or facilitated via a linking protein. STVs
and PTVs are shipped through the axon on MTs. STV and PTV transport is coordinated,
and these organelles can move together. Axonal MTs are arranged with their minus end
toward the soma, suggesting that entry into the axon requires kinesin plus-end directed
motors.
8
Motors
After STVs and PTVs are formed, they are tethered to microtubule (MT) motors
and shipped through the axon (Figure 1.1). STVs and PTVs are associated with both
kinesin and dynein motors, which move toward and away from MT plus ends,
respectively (63). Since axonal MTs are oriented with their plus ends facing distally (64-
67), kinesins drive vesicles anterogradely. STVs are transported via the kinesin-3/KIF1A
and KIF1B motors and kinesin-1/KIF5, while PTVs are transported via kinesin-1/KIF5
(Table 1) (52, 68-79). Dynein drives both PTVs and STVs towards the soma (Table 1)
(80-86). Because STVs and PTVs are likely linked to anterograde and retrograde motors
simultaneously, they typically move in a bi-directional fashion (47, 48, 50, 53, 58, 60, 73,
87, 88). This movement is frequently interrupted by brief pauses. While the exact cause
of these pauses is unknown, the sites in the axon where STVs pause are preferential sites
of presynaptic terminal formation (51). Previously, it was not known whether STV and
PTV transport is coordinated within the axon. The experiments described here in Chapter
2 attempt to determine the coordination of STV and PTV transport in order to identify a
potential novel mechanism of rapid synapse formation via the instantaneous recruitment
of SV and CAZ components to a developing synapse.
Linkage to motors
Both PTVs and STVs can attach to kinesins through linker proteins, and
regulation of this interaction is critical for synapse development (Table 1). PTVs are
primarily linked to KIF5B through syntabulin (73, 74). Syntabulin simultaneously binds
to the KIF5B heavy chain and the CAZ protein syntaxin (74). Interrupting this interaction
disrupts PTV transport in the axon and leads to a decrease in the number of functional
9
presynaptic terminals (73). A number of kinesin/cargo linking proteins enable transport
of STVs. Invertebrate studies have identified syd-2/liprin-as a link between STVs and
kinesins (77, 89, 90). Disrupting this link leads to defective axonal transport and poor
incorporation of presynaptic components into synapses (77, 89). In addition, the protein
DENN/MADD (Differentially Expressed in Normal and Neoplastic cells/MAP kinase
Activating Death Domain) mediates transport of STVs in mammalian neurons by
simultaneously binding KIF1 and the STV-associated small GTPase Rab3 (91).
DENN/MADD only binds to the GTP-associated form of Rab3 and can also function as a
Rab3 GDP-GTP exchange factor (92-94). Consistent with a broader role for GTPases in
regulation of the STV/KIF1 linkage and subsequent synapse formation, the GTPase Arl-8
is critical for proper presynaptic localization in C. elegans (95). Specifically, the GTP-
bound form of Arl-8 directly interacts with UNC-104/KIF1A to ensure that aggregation
of presynaptic components does not occur prematurely in proximal axonal sections (95,
96). Further work in C. elegans has revealed that the SV associated protein SAM-4 also
interacts with the cargo-binding domain of unc-104/KIF1A and mediates STV transport
(97). In addition, unc-104/KIF1A can directly bind to STVs through
phosphatidylinositol-4,5-biphosphate (PI(4,5)P(2)), an integral lipid component of the
STV membrane, and disrupting this interaction leads to aberrant STV trafficking (98, 99).
The physical links between presynaptic transport vesicles and dynein are less well
characterized. Dynein consists of two large heavy chains that mediate ATP hydrolysis
and MT binding. The heavy chains are dimerized by five different intermediate/light
chains. These light chains also link dynein to adaptor complexes that mediate cargo
binding (84, 100). One of these adaptor complexes, dynactin, links STVs to dynein, as
10
alterations to its p150Glued
subunit leads to defective axonal transport of SV proteins (81,
85). In contrast, PTVs can directly interact with dynein light chains via bassoon, and
disrupting this interaction leads to aberrant trafficking of PTVs and altered protein
composition at synapses (83).
Additional mechanisms for regulation of STV and PTV transport
STV and PTV transport is regulated through a variety of dynamic mechanisms
(Figure 1.2). For example, phosphorylation of kinesin reduces binding to STVs (75, 101).
In addition, post-translational modification of MTs affects motor binding to MTs (102-
105). Moreover, STV and PTV movement through the axon can be altered by physical
impediments, such as reaching the end of a MT, binding of MT-associated proteins
(MAPs), or a traffic jam caused by additional motors and cargo binding the same MT
(106, 107). Neuronal activity can also affect axonal transport, possibly through calcium
influx within the axon (51, 108). The number and types of motors on individual vesicles
can vary as well, affecting transport (109). Finally, actin-based transport via myosins may
contribute to trafficking of presynaptic components, especially within the axonal initial
segment and at sites of synaptic protein recruitment (51, 103, 110-113).
11
Figure 1.2: Mechanisms of regulation of STV and PTV transport. Trafficking of
STVs and PTVs within the axon can be modulated in a number of ways. Regulation of
the following direct interactions between motor, cargo, and/or cytoskeletal elements can
potentially alter STV and/or PTV transport: (1) kinesin attachment to MTs, (2) kinesin
phosphorylation, (3) linker molecules between kinesin and cargo, (4) binding of myosin
motors to cargo, (5) binding of myosin motors to actin and/or actin polymerization at
specific sites in the axon, (6) direct physical interaction of kinesin or dynein with cargo,
(7) dynein attachment to MTs. Indirect regulation involving a number of other factors
might also affect STV and PTV trafficking: (1) motors and cargo reaching the ends of
MTs, and/or switching to adjacent MTs, (2) MT-associated proteins (MAPs) that alter
transport, (3) increased cytosolic Ca2+
, via influx and/or intracellular sources, (4) traffic
jams with other motors and cargo (synaptic or non-synaptic) on the same MT, (5) post-
translational modifications of MTs, such as acetylation or de-tyrosination. It is not yet
clear which of these mechanisms are employed to deposit synaptic material at sites of
synapse formation.
12
For a presynaptic terminal to form, STVs and PTVs must be deposited at sites of
axo-dendritic contact, requiring local cessation of transport. While many of the
mechanisms described above regulate transport and are important for synapse formation,
it remains unclear which of these mechanisms are used locally at sites of presynaptic
terminal formation. Some of these mechanisms may contribute to synapse assembly by
globally regulating transport within the axon, essentially controlling the supply of
synaptic building materials. For instance, in C. elegans, disrupting axonal transport of
presynaptic components leads to aberrant synapse formation (95, 96), and KIF1A is
critical for synaptogenesis in the hippocampus (114). Conversely, other mechanisms may
be engaged specifically at sites of synapse assembly in order to stop transport and deposit
the building materials. It will be important to distinguish between these possibilities in
the future. This will likely require local manipulation of each of these mechanisms at
sites of presynaptic assembly combined with high-resolution time-lapse imaging.
STV and PTV recruitment into developing synapses – a dynamic process
Initial recruitment of presynaptic proteins into a developing synapse can be rapid.
PTVs and STVs can be recruited to novel synaptogenic contacts within minutes (53, 115-
117). STV and PTV co-recruitment suggests that a functional synapse can form within
this time scale (118, 119). Indeed, both STVs and PTVs can instantly become trapped at
sites of synaptogenic signaling as they are trafficked through the axon (88, 115, 116).
Because STVs can cycle with the plasma membrane and likely release neurotransmitter
prior to recruitment (47, 48, 50, 51, 54-58, 120), new sites of recruitment probably have
at least some immediate functional capability. However, STV cycling is mechanistically
and functionally different than SV cycling at mature synapses (54, 56). The time required
13
for functional maturation is unclear. Live imaging of developing synapses followed by
retrospective electron microscopy indicated that the time required to accumulate a full
complement of synaptic structures is variable and often takes hours (121, 122). In
addition, it can take hours for axonal regions exposed to a postsynaptic synaptogenic
signal to become saturated with newly formed synapses (88, 115).
Questions also remain about the order and potential coordination of STV and PTV
recruitment. Since many CAZ proteins tether SVs at mature synapses, PTVs are typically
thought to arrive at developing synapses first, where they can then recruit STVs (123). In
support of this theory, the average time it takes for PTVs to be recruited to developing
synapses tends to be shorter than the average time for STV recruitment, albeit within
minutes of each other (116, 118). However, initial recruitment of STVs and PTVs has
only been observed in separate cells (i.e. observing STV recruitment in one neuron and
PTV recruitment in a different neuron). In both of the above studies, the time-course of
recruitment for SV and CAZ proteins overlaps (116, 118). In addition, STVs and PTVs
can co-localize with each other within the axon, suggesting that an entire complement of
synaptic proteins might be recruited to a developing synapse simultaneously (49, 87).
Furthermore, clustering of the trans-synaptic adhesion protein neurexin (see below)
within the axon leads to the formation of presynaptic terminals (124, 125). Finally, live
imaging of STVs and PTVs in separate axons of zebrafish embryos, followed by
retrospective immunocytochemistry identified STVs as the first transport vesicle that
arrives at the synapse (126). Therefore, live time-lapse imaging of both STVs and PTVs
within the same axon is needed to fully understand this critical event in the formation of a
presynaptic terminal.
14
Sites of Preferential Synapse Formation
Synapses form at discrete sites within neurons and are restricted in their overall
size (41, 127). Even when synaptogenic signaling is initiated in a continuous axonal area
tens of m in length, presynaptic terminals appear at only certain points within this
region (51, 88, 115, 128, 129). In addition, sites in the axon where STVs and PTVs pause
are sites of preferential synapse development (51, 87). Pausing at these preferential sites
might act as a mechanism for localizing presynaptic proteins to these areas, allowing
rapid incorporation of STVs and PTVs into the developing synapse because these
components are already present at the site once synaptogenic signaling commences.
However, this hypothesis needs to be tested directly. Many dendritic filopodia make
transient contact with neighboring axons, with only a portion of axo-dendritic contacts
becoming stabilized (130-135). Therefore, dendritic filopodia may sample the axon to
find the suitable synaptic sites.
The molecular mechanisms responsible for creating preferred sites of synapse
formation within the axon are unknown. One possibility is that axonal synaptogenic
adhesion proteins are involved. For example, distinct patterns or states of neurexin
oligomerization might identify preferred sites of synapse formation. However, the
interaction of neurexin with neuroligin is likely not required for initial formation of these
sites, as they can be identified prior to axo-dendritic contact (51). Future studies directed
at uncovering the molecular components responsible for these preferential sites will be
critical to understand the initial processes behind presynaptic terminal formation.
Synaptogenic signaling through trans-synaptic adhesion proteins
15
For a presynaptic terminal to form, STVs and PTVs must be recruited to a
specific site in the axon in response to a synaptogenic signal. This process is triggered by
physical contact between two cells – in the brain, typically between an axon and dendrite.
When axo-dendritic contact is established, binding occurs in trans between the
extracellular domains of adhesion proteins that are localized to axons and dendrites. This,
in turn, leads to synaptogenic signaling within the axon and dendrite, subsequent
recruitment of synaptic components to both sides of the contact, and eventual formation
of a functional synapse (136).
A number of synaptogenic trans-synaptic adhesion interactions have been
identified (Table 2). Differential expression of these proteins between neuronal
populations may help organize brain circuitry by controlling the specificity of synapse
formation (136-138). For example, two isoforms of neuroligin - neuroligin-1 and
neuroligin-2 - are specifically localized to excitatory and inhibitory synapses,
respectively (139, 140). This specificity appears to be driven by the axon, arising from
the interaction preferences of presynaptic neurexins found in excitatory and inhibitory
axons (141, 142). Likewise, isoforms of SynCAM are differentially expressed in regions
of the brain and display specific binding patterns between isoforms (143, 144). Binding
specificity can be mediated by alternative splicing. The neuroligin-neurexin interaction is
extensively regulated through this mechanism, and different binding pairs exhibit
different functional properties, including differences in the rates of presynaptic terminal
formation (115, 145-152).
Trans-synaptic adhesion proteins also display redundant functions during synapse
formation. Mice lacking neuroligin-1, 2, and 3 have a normal number of synapses (153).
16
Likewise, synapse number is normal in mice lacking all isoforms of the -neurexin gene
(154). Although synaptic function is altered in these models, the ability to form synapses
remains intact. However, the relative expression of neuroligin between neighboring cells
in the brain does regulate synapse number (155). This suggests that the
neuroligin/neurexin interaction is important for synapse development in a competitive,
non-cell-autonomous manner. In contrast, increasing or knocking out SynCAM increases
or decreases hippocampal synapse number, respectively (156). While alterations in
synapse number point to a synaptogenic function of SynCAM, the magnitudes of the
changes were approximately 10-20% (156), illustrating that other synaptogenic signals
are involved in this process. In fact, there is no individual molecule that is absolutely
critical for synapse formation in the brain. Therefore, determining how the multitude of
trans-synaptic adhesion signals cooperate to control synapse formation will be critical for
a full understanding of synapse and circuit formation in the brain.
17
Table 2: Trans-synaptic adhesion partners
Presynaptic Postsynaptic References
Homophilic
SynCAM
SynCAM (129, 156, 157)
N-Cadherin
N-Cadherin (158-161)
Amyloid Precursor Protein
(APP)
Amyloid precursor protein
(APP)
(162)
Heterophilic
Neurexin Neuroligin (124, 128, 139, 163, 164)
Neurexin Leucine Rich Repeat
Transmembrane Protein 2
(LRRTM2)
(165, 166)
Neurexin
GluR2 + Cerebellin 1
Precursor Protein
(167)
Protein Tyrosine
Phosphatase Receptor T
(PRPRT)
Neuroligin (168)
Protein Tyrosine
Phosphatase Sigma (PTP)
TrkC (169)
Protein Tyrosine
Phosphatase Delta (PTP)
Interleukin-1-receptor
accessory protein like 1
(IL1RAP)
(170-172)
Ephrin-B EphB (173, 174)
18
Latrophilin 1
Lasso/Teneurin-2 (175)
Netrin-G1, Netrin-G2,
Leukocyte Common
Antigen-Related Protein
(LAR)
Netrin-G Ligands (NGL-
1,2,3)
(176-178)
19
Signaling downstream of trans-synaptic adhesion
The downstream signaling pathways generated through trans-synaptic adhesion
are less clear, especially presynaptically. The interaction between neurexin and neuroligin
is the best characterized of the trans-synaptic proteins (Figure 1.3), and many of the
presynaptic signaling mechanisms that act down-stream of neurexin are likely to be
involved in synapse assembly induced by other trans-synaptic interactions.
The MAGUK protein CASK can bind to neurexin (179). CASK, in turn, binds to
syndecan-2 and protein 4.1 (180, 181). Synaptic proteins might be recruited to this
complex either through actin-mediated recruitment via filamentous-actin (F-actin)
nucleation on protein 4.1 or through phosphorylation of various substrates by CASK
itself (182, 183). Mint1 also binds to neurexin and can link neurexin, CASK, Veli/MALS,
and the active zone protein Munc-18 into a single complex (184, 185). This
CASK/Mint1/Veli complex may also play a role in SynCAM and APP mediated
presynaptic development (129, 162, 186). Furthermore, the tandem-PDZ protein syntenin
can directly interact with neurexin and is also linked to the CASK/Mint2/Veli complex
through syndecan-2 (187, 188). Syntenin also interacts with the active zone protein
CAST1/ERC2/ELKS via its PDZ domain, which in turn, binds RIM1, liprin-, bassoon,
and piccolo (188-192). Piccolo facilitates F-actin formation, which can lead to
recruitment of additional synaptic components (193), while bassoon interacts directly
with dynein to regulate recruitment of CAZ components (83).
Liprin- interacts with a number of these complexes, and disrupting this
interaction leads to aberrant synapse formation (194). Liprin- (syd-2 in C. elegans) is
20
involved in active zone morphology and recruitment of active zone proteins (195-201).
Interestingly, mutations that inhibit liprin- binding to the CASK/Mint1/Veli complex
have been linked to X-linked mental retardation (202). Because liprin- also links STVs
to KIF1, these interactions might be critical for initial recruitment of STVs into the
synapse (77, 89, 97, 203). Liprin-α acts downstream of syd-1 during presynaptic
development in C. elegans (196, 197). In Drosophila, Syd-1 binds to neurexin through its
PDZ domain and mediates neurexin clustering within the axon (125), which appears to be
necessary for neurexin’s synaptogenic function (88, 124, 204, 205). Recently,
mammalian syd-1 was shown to organize synapses through binding with other active
zone proteins, including liprin-2 and munc18-1 (206). However, it is unclear if
mammalian syd-1 binds to neurexin, as the PDZ domain that is critical for neurexin/syd-1
binding in Drosophila is absent in mammals.
21
Figure 1.3: Recruitment of presynaptic cargo downstream of neurexin-dependent
synaptogenic adhesion. Neurexin interacts with a variety of presynaptic proteins;
however, the mechanisms through which STVs and PTVs are incorporated into the
22
synapse remain unclear. This figure illustrates potential mechanisms that might be
important for this process. (1) Bassoon directly interacts with dynein light chains 1 and 2,
as well as myosin V. Blocking the bassoon/dynein interaction leads to aberrant bassoon
and piccolo synaptic localization. (2) The PDZ-domains of Mint1, Veli/MALS, or CASK
might interact with other synaptic proteins in STVs or PTVs in order to trap them at a
developing synapse. (3) F-actin nucleation mediated by Protein 4.1 might enable the
removal of STVs and/or PTVs from their MT highways via myosin motor binding. (4)
Liprin- can serve as a linker molecule between STVs and the KIF1 motor. Delivery of
liprin- to the synapse might serve to cluster and/or recruit other synaptic components.
Alternatively, modulation of liprin- to reduce its binding to KIF1 might play a role in
cargo delivery. (5) CASK can directly phosphorylate substrates at presynaptic terminals.
Therefore, it might phosphorylate a motor-binding protein integral to STVs or PTVs, a
motor-cargo adaptor or the motor itself, which has been shown to reduce binding between
kinesins and STVs. (6) Although neurexin interacts with a plethora of cytosolic targets,
its cytoplasmic domain is completely dispensable for many of its synaptogenic properties.
This suggests that cis interactions between the extracellular domain of neurexin and a
yet-to-be-determined trans-membrane effector molecule(s) can induce recruitment of
presynaptic proteins. The number and type of downstream recruitment mechanisms
mediated by this potential interaction is unknown.
23
Many additional aspects of trans-synaptic signaling remain unclear. For example,
trans-synaptic signaling does not affect all STVs and PTVs equally: many vesicles can
pass directly through regions of trans-synaptic adhesion without becoming recruited into
a synapse (88, 96). It was also previously not known whether STVs and PTVs were
actively transported to sites of synaptogenic signaling or passively trapped at these sites.
Furthermore it is currently unknown how trans-synaptic adhesion causes STVs and PTVs
to dissociate from their MT motors and become incorporated into the synapse.
Identifying these mechanisms and determining how they are coordinated during
synaptogenesis will be critical to understanding how presynaptic terminals form. To that
extent, experiments described here in Chapters 3 and 4 attempt to elucidate some of these
mechanisms
In this thesis, a number of major issues concerning presynaptic development will
be addressed. In Chapter 2, the trafficking of two transport vesicles carrying distinct sets
of presynaptic proteins was found to be coordinated prior to synapse formation,
uncovering a potential mechanism for rapid recruitment of components into a developing
synapse. The experiments in Chapter 3 determine the dynamics through which
presynaptic proteins arrive at developing synapses and describe potential mechanisms
through which these components become recruited. Finally, in Chapter 4, in vivo imaging
via two-photon microscopy through cranial windows is utilized to determine the
dynamics of presynaptic terminals in the developing mouse brain.
24
Chapter 2: Coordinated trafficking of synaptic vesicle and active zone proteins
prior to synapse formation
Luke A.D. Bury1
and Shasta L. Sabo1,2
Departments of 1Pharmacology and
2Neuroscience, Case Western Reserve University
School of Medicine, Cleveland, Ohio.
*Parts of this chapter were published in the journal Neural Development, Volume 6,
Issue 24, May 10, 2011.
25
Abstract
The proteins required for synaptic transmission are rapidly assembled at nascent
synapses, but the mechanisms through which these proteins are delivered to developing
presynaptic terminals are not understood. Prior to synapse formation, active zone
proteins and synaptic vesicle proteins are transported along axons in distinct organelles
referred to as piccolo-bassoon transport vesicles (PTVs) and synaptic vesicle protein
transport vesicles (STVs), respectively. Although both PTVs and STVs are recruited to
the same site in the axon, often within minutes of axo-dendritic contact, it is not known
whether or how PTV and STV trafficking is coordinated before synapse formation. Here,
using time-lapse confocal imaging of the dynamics of PTVs and STVs in the same axon,
we show that vesicle trafficking is coordinated through at least two mechanisms. First, a
significant proportion of STVs and PTVs are transported together before forming a stable
terminal. Second, individual PTVs and STVs share pause sites within the axon.
Importantly, for both STVs and PTVs, encountering the other type of vesicle increases
their propensity to pause. To determine if PTV-STV interactions are important for
pausing, PTV density was reduced in axons by expression of a dominant negative
construct corresponding to the syntaxin binding domain of syntabulin, which links PTVs
with their KIF5B motor. This reduction in PTVs had a minimal effect on STV pausing
and movement, suggesting that an interaction between STVs and PTVs is not responsible
for enhancing STV pausing. Our results indicate that trafficking of STVs and PTVs is
coordinated even prior to synapse development. This novel coordination of transport and
pausing might provide mechanisms through which all of the components of a presynaptic
terminal can be rapidly accumulated at sites of synapse formation.
26
Introduction
In the cortex, the bulk of synapse formation occurs at high rates over a period of
several weeks during early postnatal development. Formation of individual synapses is
triggered when axons and dendrites contact one another (45, 46). For these axo-dendritic
contacts to stabilize and form a synapse, synaptic vesicle and active zone proteins must
be recruited to the site of contact very rapidly, within minutes to hours (51, 53, 115, 117,
118, 207-209). This rapid assembly is remarkable, considering that each presynaptic
protein needs to be transported from the soma to individual sites of synapse formation
along the axon.
Recent work using live imaging of developing neurons has revealed mechanisms
that might facilitate rapid synapse assembly. For example, rather than transport each
molecule individually, neurons package multiple presynaptic components into vesicles in
the cell body and transport the entire group together. Two major groups of proteins are
transported in this fashion: active zone proteins are transported in piccolo-bassoon
transport vesicles (PTVs), while synaptic vesicle-associated proteins are transported in
synaptic vesicle protein transport vesicles (STVs) (47-51, 53, 59, 60, 210). PTVs and
STVs can be distinguished both morphologically and biochemically. PTVs have been
described as dense-core vesicles or aggregates of vesicles and proteins that range in size
from approximately 80nm in diameter for dense core vesicles to 130nm by 220nm in area
for aggregates (49, 59). Proteins transported by PTVs include piccolo, bassoon, N-
cadherin and syntaxin (49, 59, 60, 73, 74). STVs are heterogeneous in size and shape and
are comprised of both tubulovesicular and clear core vesicles (47, 48, 53). Proteins
carried by STVs include synaptophysin, synapsin Ia, synaptotagmin and
27
synaptobrevin/vesicle-associated membrane protein 2 (VAMP2) (47-49, 51, 53). Both
types of vesicles are packaged via the trans-Golgi network before being transported
through the axon (48, 49, 61).
Rapid assembly of presynaptic terminals can also be facilitated by having the
source of synaptic proteins nearby and readily available. Indeed, many STVs and PTVs
are mobile within axons well before synapse formation. STVs and PTVs travel in a
saltatory fashion in both the anterograde and retrograde directions along the entire length
of the axon [4, 8, 11, 13-15, 17, 19]. This provides a readily-available pool of synaptic
vesicle and active zone proteins wherever and whenever axo-dendritic contact occurs.
Since both STVs and PTVs must be delivered to the same site during synapse
assembly, we hypothesized that trafficking of STVs and PTVs might be coordinated even
prior to synapse assembly, providing an additional mechanism to facilitate rapid
accumulation of the full complement of proteins required for synaptic transmission. Such
coordination could arise through co-transport of STVs and PTVs, through pausing of
STVs and PTVs at the same sites, or both. Coordinated pausing presents a particularly
intriguing possibility since sites along the axon where STVs pause their transport are
preferential sites of synapse formation (51). It was previously suggested that cues that
control pausing at these sites could promote rapid synapse assembly by increasing the
probability that STVs are at or near any given site when axo-dendritic contact occurs. If
this model is correct, then synapse assembly would be most efficient if PTVs are also
attracted to these same sites. However, it is not yet known whether PTVs also pause at
these sites and, if so, whether they do so simultaneous with STVs. A recent report
demonstrated that PTVs and STVs (particularly those resembling small, clear vesicles)
28
can be observed tethered together in electron micrographs of developing axons (49).
These aggregates could correspond to PTVs and STVs that are either being co-
transported or pausing at the same site at the same time.
Here, we tested whether transport and pausing of PTVs and STVs is coordinated
prior to synapse assembly using time-lapse confocal imaging of green and red fluorescent
protein-tagged synaptic vesicle and active zone proteins within the same axon. We found
that a significant portion of STVs and PTVs move together and share pause sites.
Interestingly, both STVs and PTVs preferentially paused at these sites when another
vesicle was present. These observations raised the question of whether a direct interaction
between STVs and PTVs coordinates their transport and pausing. Reducing PTVs in the
axon minimally affected the movement and pausing of STVs, arguing against this
mechanism and suggesting that other unidentified signals are responsible for STV and
PTV coordination. These findings represent novel mechanisms that can facilitate the
rapid recruitment of presynaptic proteins to the same site within the axon and, therefore,
promote synapse development.
29
Results
STVs and PTVs can move together
STVs and PTVs move in a similar saltatory fashion within the axon in both the
anterograde and retrograde directions (47, 48, 51, 53, 58, 60, 73). However, because
STVs and PTVs have almost always been imaged separately, it is not yet known if and
how STV and PTV trafficking interrelate. To determine this, we used time-lapse imaging
of neurons co-transfected with fluorescent STV and PTV markers. In this assay, 5-6 DIV
rat cortical neurons were co-transfected with GFP-bassoon to label PTVs and either
synaptophysin-mcherry or synaptophysin-mRFP to label STVs. Previous studies have
indicated that fluorescent proteins attached to bassoon and synaptophysin effectively
label PTVs and STVs, respectively (48, 50, 53, 60, 61, 115, 210, 211). 1-2 days after
transfection, time-lapse images of the fluorescently labeled vesicles were obtained from
individual axons. Images were taken every 10s for 7.5 minutes.
From the kymographs in Figure 2.1A, it is clear that PTVs and STVs moved
together in both anterograde and retrograde directions. To quantify these correlated
movements, the positions of individual STV and PTV puncta were tracked independently
while blind to the other channel, and then their movements were compared. To
determine whether these correlated movements could be accounted for by chance, the
experimental data were ultimately compared to model axons where the initial positions of
all of the puncta were randomized but the density of puncta and properties of their
movements were derived from the experimental data (Figure 2.1B).
30
During the imaging period, 18.3% of PTVs moved with STVs (n = 224 vesicles),
while 37.1% of STVs moved with PTVs (n = 97 vesicles; Figure 2.1C). STVs and PTVs
that moved together also spent the majority of their time together (Figure 2.1D). The
prevalence of correlated STV and PTV movements was over 10-fold higher for the
experimental data than predicted by the same analysis of model axons (Figure 2.1C;
model: n = 22400 PTVs and 9700 STVs). In addition, the percentage of time STVs and
PTVs spent moving together was over 2.5-fold higher expected by chance (Figure 2.1D).
These data indicate that a population of STVs and PTVs travel together within the axon,
which could allow both sets of presynaptic proteins to be distributed together to sites of
synapse formation.
31
Figure 2.1: STVs and PTVs move together. (A) Kymographs of an axon segment
showing the movements of PTVs (GFP-bassoon; green) and STVs (synaptophysin-
mcherry; magenta) in the same axon. Yellow lines underline the moving STVs and PTVs.
Bottom panel, overlay of STV and PTV fluorescence. On the ordinate axis, one pixel
corresponds to 10s. On the abscissa, the scale bar corresponds to 10µm. (B) Simulations
of model axons were performed by randomizing initial positions of vesicles while
maintaining movement and pausing characteristics of the original imaged vesicles.
Diagrams illustrate kymographs of 3 simulations of model axons. Movements of two
32
vesicles are shown in each model axon. (C) Plot showing the percentage of PTVs that
move with STVs (green) and STVs that move with PTVs (magenta). PTV and STV
simulations (light green and magenta, respectively) correspond to the predicted values
from the simulations. (D) Quantification of the percentage of time that PTVs and STVs
spent together. Both the percentage of vesicles that moved together and the time spent
together were substantially larger in the data than is predicted by chance (simulations).
33
STVs and PTVs display similar pausing characteristics
Although the movements of both STVs and PTVs have been extensively
described individually, the pauses between movements have been less thoroughly
analyzed and have not been compared. Comparing STV and PTV pausing behavior is
important since STVs preferentially pause at sites of eventual synapse formation (51, 53).
When PTV behavior was examined, it was clear that PTV pausing exhibited many of the
properties previously described for STVs (Figure 2.2A-B and (51)). For example, the
same PTV could be observed repeatedly pausing at a given site (Figure 2.2B, top panel).
In addition, multiple PTVs paused at the same site, both sequentially and simultaneously
(Figure 2.2B, middle and bottom panels, respectively). When STV movements were
quantified, the mean STV pause frequency was 0.0081 +/- 0.0002 pauses/second (n = 262
vesicles, Figure 2.2C). The average STV pause duration was 107.4 +/- 3.4 seconds (n =
954 pauses, Figure 2.2D). Both measurements concur with previous observations (51).
As shown in Figure 2.2C-D, PTV pause frequency and duration in 7-8 DIV neurons were
nearly identical to the STV data, with PTVs pausing at a frequency of 0.0082 +/- 0.0003
pauses/second and average duration of 108.3 +/- 3.5 seconds (n = 253 vesicles, n = 933
pauses). The similarities between the pausing behavior of STVs and PTVs raise the
question of whether PTVs might pause with STVs at sites of eventual synapse formation.
Like STVs (51), regulation of PTV pausing could be an important target of signals that
control synapse assembly.
34
Figure 2.2: PTV pausing is qualitatively and quantitatively similar to STV pausing.
(A) Time-lapse images of PTVs labeled with GFP-bassoon. Individual PTVs are tracked
with red, yellow or blue arrows. Most PTVs paused, and pauses were of varied durations.
Frames were collected at the indicated times. Scale bars, 5 µm. (B) Kymographs
35
demonstrating the movements of PTVs in 3 axons. Individual PTVs paused repeatedly at
the same site (top panel), and multiple PTVs paused at the same site (middle and bottom
panels). Pausing of multiple PTVs at a given site occurred both sequentially (middle
panel) and simultaneously (bottom panel). Individual vesicles are highlighted in different
colors for visualization. Pause sites are outlined in orange. On the ordinate axis, one pixel
corresponds to 10s. (C-D) STVs (magenta) and PTVs (green) paused at similar
frequencies (C) and for similar mean durations (D). Data represent the mean + S.E.
36
STVs and PTVs pause simultaneously at the same sites in the axon
For a synapse to develop, both active zone and synaptic vesicle proteins need to
be recruited to the same site. Sharing a pause site provides a potential mechanism for
recruiting both STVs and PTVs to the same spot in the axon. Since places where STVs
pause are preferred sites of synapse formation, it seemed likely that PTVs also pause at
these sites. To test this hypothesis, STVs and PTVs were imaged and analyzed as
described above. From the time-lapse panels and kymographs shown in Figure 2.3A-B, it
is clear that both types of vesicles paused at sites where the other type of vesicle also
paused.
To determine whether pausing of STVs and PTVs at the same sites was
significantly greater than would be predicted by chance, pausing at the same sites was
quantified in each axon then compared to simulations in model axons (Figure 2.3C). For
STVs, 47.9% of pauses were at sites where a PTV also paused (n = 305 pauses, Figure
2.3D), compared to 24.5% using the same analysis of model axons (n = 33967 pauses,
Figure 2.3D). This indicates that STVs preferentially pause at PTV pause sites.
Likewise, 24.5% of all PTV pauses were at sites where an STV also paused (n = 620
pauses, Figure 2.3D), compared to 14.0% using the same analysis of model axons (n =
69263 pauses, Figure 2.3D), indicating that PTVs preferentially pause at STV pause sites.
Some STVs do not encounter PTV sites during our imaging time window and vice versa.
To account for this, we also quantified pausing at the same site with the analysis limited
to STVs and PTVs that had the opportunity to pause at PTV and STV sites, respectively
(Figure 2.3E). 57.8% of STVs that encountered a PTV site paused (n = 135), and 84.4%
of PTVs that encountered STV sites paused at those sites (n = 90). PTVs are slightly
37
more likely to pause at STV sites; however, this difference appears to be an inherent
property of the vesicles and is accounted for in model axons (see Figure 2.4A,C). It has
been shown previously that STV pause sites are preferred sites of synapse formation, and
PTVs pause at these same sites; therefore, PTVs pause at sites of synapse formation.
38
Figure 2.3: STVs and PTVs pause at the same sites. (A) Time-lapse images of a
segment of axon expressing both synaptophysin-mcherry (magenta) and GFP-bassoon
39
(green). In each panel, fluorescence signals from the two channels are overlaid, and
colocalization is indicated by white. Orange box, site where a PTV is paused. The arrow
tracks a STV that then pauses at that site. Scale bars, 5 µm. (B) Kymographs showing
two examples of STVs and PTVs that pause at the same sites. On the ordinate axis, one
pixel corresponds to 10s. Bottom panel, overlay of STV and PTV fluorescence. Orange
boxes outline 3 shared pause sites. (C) Simulations of model axons were performed by
randomizing initial positions of vesicles while maintaining movement and pausing
characteristics observed for the original experimental vesicles. Diagrams show the
superimposed locations of all pause sites for individual STVs and PTVs in model axons.
Three simulations of the same experimental data are shown. STV pause sites are
indicated with a dot while PTV pause sites are indicated with a “+” sign. Each color
represents an individual vesicle. Cyan and magenta boxes outline the pause sites of the
same PTV and STV, respectively, in each model axon. For each axon imaged, 100 such
simulations were performed, allowing estimation of the degree of co-transport and co-
pausing expected from chance alone. (D) Plot illustrating the percentage of PTV (green)
and STV (magenta) pause sites that are shared with STVs and PTVs, respectively. The
fraction of shared sites is much higher than predicted by chance (via simulations, light
green and light magenta). (E) A large majority of PTVs that encountered STV pause
sites paused at those sites. Similarly, most STVs that encountered PTV pause sites then
paused at those sites. Error bars display the 95% confidence interval.
40
Since both PTVs and STVs pause in the same places, the next question addressed
was whether STVs and PTVs stop at these sites at the same time. Synapse assembly is a
rapid process, and concurrent attraction of both STVs and PTVs could provide a
mechanism for efficient assembly of presynaptic terminals. To determine whether PTVs
and STVs are simultaneously attracted to sites of synapse formation, we quantified
whether STVs preferentially pause at PTV pause sites when a PTV is present. As shown
in Figure 2.4, STVs were more likely to pause at a site that contained a PTV than a site
that did not. When STVs encountered pause sites with PTVs present, 76.4 +/- 11.2% of
STVs paused with the PTV (n = 55; Figure 2.4A). In contrast, only 45.0 +/- 10.9% of
STVs paused at PTV pause sites when a PTV was not there (n = 80). This 1.7-fold
difference in the likelihood of pausing with a PTV could not be accounted for by chance
because there was no interaction between STVs and PTVs when the same analysis was
performed on data obtained from simulations with model axons (Figure 2.4A). In the
model, 52.4 +/- 1.9% of STVs paused at PTV pause sites with a PTV present (n = 2334),
and 51.6 +/- 1.3% of STVs paused at PTV pause sites without PTVs present (n = 5767).
The pause duration of an STV stopped at a PTV pause site was also measured. In
contrast to the likelihood of pausing, the pause duration of STVs is not affected by the
presence or absence of PTVs (Figure 2.4B). The average pause duration for STVs
paused at sites with PTVs present was 101.0 +/- 13.4 seconds (n = 42 pauses), while the
average pause duration for an STV paused at a site without a PTV present was 94.2 +/-
20.1 seconds (n = 36).
It is not yet known whether STVs and PTVs must be recruited to nascent
presynaptic terminals in a defined order, so we also tested whether PTVs are more likely
41
to pause at sites that contain STVs. As shown in Figure 2.4C, PTVs paused at sites that
contained STVs at a significantly higher rate (93.6 +/- 7.0%, n = 47) than at sites where
STVs had previously paused but were no longer present (74.4 +/- 13.0%, n = 43). In
contrast, this effect of STVs on PTV pausing behavior was not seen with analysis of the
randomized model: in the simulations, 76.0 +/- 2.0% of PTVs paused at STV sites with
an STV present (n = 1591), and 74.7 +/- 1.1% of PTVs paused at STV sites without
STVs present (n = 4906). There was no difference in the probability that either vesicle
arrived first at a shared pause site, based on 95% CI (data not shown). Similar to STVs
pausing at PTV sites, the mean pause duration of PTVs pausing at STV pause sites is not
affected by the presence of STVs (Figure 2.4D). The average pause duration for PTVs
paused at sites with STVs present was 100.5 +/- 12.4 seconds (n = 44 pauses), while the
average pause duration for PTVs paused at sites without STVs present was 80.9 +/- 15.3
seconds (n = 32 pauses). These data demonstrate that PTVs prefer to pause at sites that
contain STVs and vice versa, suggesting that STVs and PTVs are attracted to the same
sites at the same time.
42
Figure 2.4: STVs and PTVs preferentially pause at the same sites at the same time.
(A) STVs (magenta) are significantly more likely to stop at a PTV pause site when PTVs
are at the site. This preference cannot be accounted for by chance since no dependence on
the presence of PTVs was seen in simulations (light magenta). The data presented
correspond to the mean + 95% confidence intervals. *, 95% confidence intervals do not
43
overlap. (B) STVs pause for similar lengths of time regardless of whether a PTV is
present at the pause site. Data are presented as mean + S.E. (C) PTVs (green) are more
likely to pause at a site when an STV is present. The same dependence was not observed
in simulations (light green). (D) PTVs pause for similar lengths of time, regardless of
whether STVs are present at the pause sites. Data are the mean + S.E.
44
Recruitment of a PTV enhances accumulation of additional PTVs at sites of synapse
formation
Assembly of presynaptic terminals involves recruitment of multiple PTVs (60).
Therefore, we wondered whether PTVs might also increase attraction of other PTVs to
sites of synapse formation. To test this, we quantified the percentage of PTVs that
paused at sites where another PTV was either present or had previously paused. When a
PTV was already anchored at a particular pause site, there was high likelihood of an
additional PTV pausing at that site: 83.3 +/- 13.3% of PTVs paused when they
encountered sites with other PTVs (n = 30; Figure 2.5A). However, when a PTV was not
present at the pause site, the likelihood that a PTV would pause was significantly lower
(42.5 +/- 15.3%, n = 40). This nearly 2-fold increase in attraction of PTVs to sites
containing other PTVs could not be explained by chance since the increase was not
observed in our simulations. In model axons, PTVs paused at 67.1 +/- 1.6% of sites with
other PTVs (n = 1267) and 68.1 +/- 1.2% of sites without other PTVs (n = 5827; Figure
2.5A). This indicates that the presence of a PTV at a pause site promotes recruitment of
additional PTVs to that site.
Interestingly, the presence of an STV at a particular pause site did not
significantly increase the probability of an additional STV pausing at that site (chance of
pausing with another STV present = 71.4 +/- 23.7%, n = 14; without another STV present
= 60.5 +/- 15.5%, n = 38; Figure 2.5B). As expected, no dependence of STV pausing on
the presence of other STVs was observed in our randomized model axon (STV present =
56.8 +/- 2.4%, n = 1665; STV absent = 59.8 +/- 1.6%, n = 3443; Figure 2.5B). These data
45
suggest that STVs do not interact with one another in a way that promotes recruitment to
sites of synapse formation.
46
Figure 2.5: Multiple PTVs are attracted to the same sites. (A) PTVs are more likely to
pause when other PTVs are present (green). Spatially randomized simulations are shown
in light green and cannot account for the tendency of PTVs to pause simultaneously with
other PTVs. (B) STVs are not more likely to pause when other STVs are already present
(magenta). Simulations are shown for comparison (light magenta). Error bars represent
95% confidence intervals. *, 95% confidence intervals do not overlap.
47
STV pausing is only mildly dependent on PTVs
The data presented above demonstrate that pausing of STVs at sites of synapse
formation is enhanced at sites that contain PTVs and vice versa. This raises the question
of whether a direct physical interaction between STVs and PTVs is important for
recruitment of synaptic proteins to sites of synapse formation. If so, then STV pausing
should be altered if PTVs are decreased or eliminated from the axon. Conveniently, a
method for disrupting PTV transport – and, therefore, decreasing PTV density in the axon
-- has recently been described (73, 74). This approach utilizes a dominant negative
construct that disrupts the connection between the PTV-associated protein syntaxin and
syntabulin, which links PTVs to the kinesin motor KIF5B. This construct mimics the
syntaxin binding domain (SBD) of syntabulin and interferes with the syntaxin-syntabulin
interaction. Expression of syntabulin-SBD has been shown to specifically prevent the
majority of PTVs from being transported out of the cell body and into the axon without
directly affecting STV transport or other KIF5 cargo (Figure 2.6A; (73, 74)). Imaging of
our cortical cultures confirmed that the density of endogenous piccolo puncta is
dramatically decreased in axons of neurons expressing syntabulin-SBD fused to GFP
when compared to neurons expressing GFP alone (Figure 2.6B).
Consistent with the published data, transfection of neurons with the syntabulin-
SBD construct did not interfere with STV transport in axons (Figure 2.6C): STVs still
moved, and the mean distance moved over the course of imaging was unchanged (No
SBD = 8.90 +/- 0.93 µm, n = 163 vesicles; SBD = 9.30 +/- 0.82 µm, n = 153 vesicles, p =
0.68). When STV pausing was quantified, expression of the syntabulin-SBD construct
yielded no change in the pause frequency (No SBD = 0.0085 +/- 0.0003 pauses/second, n
48
= 163 vesicles; SBD = 0.0080 +/- 0.0004 pauses/second, n = 153 vesicles; p = 0.62;
Figure 2.6D) but did cause a slight decrease in the average duration of STV pauses (No
SBD = 109.9 +/- 4.4 s, n = 586 pauses; SBD = 102.4 +/- 4.3 s, n = 582 pauses; p = 0.03;
Figure 2.6E). In addition, the instantaneous velocities of STVs were slightly reduced in
SBD-transfected neurons (No SBD = 0.208 +/- 0.004 µm/s, n = 729 movements; SBD =
0.193 +/- 0.004 µm/s, n = 774 movements; p = 0.003; Figure 2.6F). Although there was a
decrease in STV pause duration when PTV transport was disrupted, the magnitude of the
effect and the lack of a change in the probability of pausing argue against PTVs
themselves controlling STV pausing. These data suggest that a direct interaction between
STVs and PTVs is not responsible for the increased attraction of STVs to sites that
contain PTVs.
49
50
Figure 2.6: A direct interaction between STVs and PTVs cannot account for the
attraction of STVs to pause sites that contain PTVs. (A) PTV transport was inhibited
using dominant-negative syntabulin (syntaxin binding domain, SBD) fused to GFP. (B)
Expression of syntabulin-SBD-GFP decreases the density of PTV puncta in axons when
compared to axons expressing GFP alone. PTVs were identified by immunofluorescent
labeling for endogenous piccolo. (C) Images and kymograph (bottom) showing that STVs
(magenta) move and pause in SBD-expressing axons (green). (D) The frequency of STV
pausing was unchanged in the presence of SBD-GFP. (E) STVs pause durations are
shorter when PTV localization in the axon is disrupted. p-values are from Wilcoxon rank-
sum test. The change in pausing upon expression of SBD-GFP is not sufficient to account
for the coordinated pausing of STVs and PTVs. (F) The instantaneous velocities of STVs
are increased in SBD-expressing axons. Black arrow, mean instantaneous velocity.
51
Materials and Methods
All studies were conducted with an approved protocol from the Case Western
Reserve University Institutional Animal Care and Use Committee, in compliance with the
National Institutes of Health guidelines for the care and use of experimental animals.
Neuronal cultures and transfection
Primary neuronal cultures were prepared from postnatal rat visual cortex
essentially as described previously [4, 5, 15], except neurons were maintained in
Neurobasal-A medium supplemented with glutamax and B27 (Invitrogen). Neurons were
transfected with Lipofectamine 2000 (Invitrogen, Carlsbad, CA) 24-48 h before live
imaging. Excluding GFP-bassoon transfection, 1µg of DNA construct was combined
with 1µg of Lipofectamine 2000 in 50ul of Optimem (Invitrogen) for each 18 mm
coverslip. Transfection of GFP-bassoon was conducted in the same manner except 2µg
of DNA were used to account for its large size. With double transfection, localization of
each protein appeared similar when expressed alone. Also, the distribution and
movement of STVs labeled with synaptophysin-mcherry, synaptophysin-mRFP, and
synaptophysin-GFP all appeared similar to one another. GFP -bassoon (GFP-Bsn 95-
3938), synaptophysin-mcherry, synaptophysin-mRFP, synaptophysin-GFP and syntaxin-
SBD-GFP were generous gifts of Drs. Thomas Dresbach (University of Heidelberg,
Heidelberg), Matthijs Verhage (Vrije Universiteit, Amsterdam), Jurgen Kingauf
(University of Muenster, Muenster), Jane Sullivan (University of Washington, Seattle)
and Zu-Hang Sheng (National Institute of Neurological Disorders and Stroke, Bethesda).
52
Each of these constructs has been shown previously to be functional and properly
localized (50, 61, 73, 211-213).
Live imaging
Neurons were imaged at 7-8 DIV with a C1 Plus confocal system with a Nikon
Eclipse Ti-E microscope using a 40x Nikon Plan Apo 0.95 numerical aperture objective.
Lasers were 488 nm argon and 543 nm helium-neon. Detection filters were 515/30 nm
bandpass for GFP and 590/50 nm bandpass for mcherry/mRFP. Images were collected
every 10s, with scan times no greater than 3.3s. This imaging interval was selected as a
compromise between having a high temporal resolution and minimizing the time the
neurons were exposed to the laser to avoid any toxicity and photobleaching. A total of 45
images were collected for each time-lapse series. Imaging was conducted with constant
perfusion with artificial CSF (ACSF) (120 mM NaCl, 3 mM KCl, 2 mM CaCl2, 2 mM
MgCl2, 30 mM D-glucose, 20 mM HEPES, and 0.2% sorbitol, pH 7.3). ACSF perfusion
was performed at 25oC since STV transport and pausing are not significantly different at
ambient and physiological temperatures (51). Axons were identified using morphological
criteria, as described previously (50, 51). For dual-color imaging, channels were
collected sequentially to eliminate bleed-through and neurons were imaged in which
expression levels of both fusion proteins appeared comparable.
Immunofluorescence
Neurons were fixed for 15 minutes in 4% paraformaldahyde in PBS containing
4% sucrose, permeabilized for 5 minutes with 0.2% Triton X-100, and blocked with 10%
horse serum. The primary antibody was piccolo (Synaptic Systems), and the secondary
53
antibody was Alexa 633-conjugated goat anti-rabbit (Invitrogen). Coverslips were
mounted in Fluoromount (Fisher Scientific, Pittsburgh, PA) containing DABCO (1,4-
diazabicyclo[2.2.2]octane) (Sigma, St. Louis, MO).
Analysis and statistics
STV and PTV movements were tracked using ImageJ (NIH, Bethesda, MD). To
restrict the quantification of time-lapse movies to healthy neurons, axons were included
in the analysis only if a least 1 vesicle moved within the field of view. Regions of axon
with an intermediate STV or PTV density were imaged to allow us to track each vesicle
reliably. STV and PTV movements were tracked independently while blind to the other
channel.
For quantification of movement and pausing, positional data were imported into
Matlab and analyzed with custom-written programs (Mathworks, Natick, MA). For
pause analysis, a pause was defined as a period greater than or equal to 10s, during which
the velocity of the vesicle went to 0+/- 0.1 µm/s. The cutoff zero velocity (0.1µm/s) was
chosen based on the average diameter of vesicles, which was approximately 1µm.
Vesicles that never moved were not included in the analysis. Therefore, vesicles which
pause for very long durations might be underrepresented. Also, only vesicles that could
be tracked for the entire imaging duration were included in the analysis. Vesicles that
move at high velocities are more likely to leave the imaging area before the end of the
movie. Therefore, these vesicles might also be underrepresented in the analysis.
Given the pixel density, scan speed and average vesicle size, a vesicle was
typically imaged in approximately 10-50 ms, permitting reliable tracking of vesicles
54
based on their apparent size, shape, and intensity, with relatively low influence of vesicle
movements on these parameters. The fastest STV movements, at a maximum
approaching 1 µm/s (comparable with the fastest velocities that have been reported for
STVs (51, 53, 210)), resulted in movements of 10 µm during the imaging interval and
were easily measured. Maximal velocities of PTVs are lower than maximal STV
velocities (60, 73) and were also easily recorded.
Data are presented cumulatively with 95% confidence intervals for binomially
distributed data or as the mean +/- standard error of the mean (S.E.) where appropriate.
Confidence intervals were calculated based on a normal approximation, and data sets
were considered significantly different if their 95% confidence intervals were non-
overlapping. For data presented as means, significance was evaluated using the Wilcoxon
rank sum test.
Randomized model
In some cases, experimental data were compared to randomized simulations of
vesicle movement. For each axon that was imaged, a model axon was generated with the
same number of vesicles and axonal length as each experimental axon. In these models,
the initial position of each vesicle was randomized. However, the number, timing and
direction of movements, as well as the instantaneous velocity, and pause duration of each
vesicle remained the same. The newly generated time-lapse series were then analyzed in
the same manner as the experimental data. Each simulation was performed 100 times per
axon to enhance statistical analysis.
55
Acknowledgements
We thank Michael Sceniak for advice and help with Matlab programming, for help with
design of the simulations and for helpful discussions about the project. We also thank
Corbett Berry and Louie Zhou for technical assistance. We are grateful to Drs. Thomas
Dresbach (University of Heidelberg, Heidelberg), Matthijs Verhage (Vrije Universiteit,
Amsterdam), Jurgen Kingauf (University of Muenster, Muenster), Jane Sullivan
(University of Washington, Seattle) and Zu-Hang Sheng (National Institute of
Neurological Disorders and Stroke, Bethesda) for GFP-bassoon (GFP-Bsn 95-3938),
synaptophysin-mcherry, synaptophysin-mRFP, synaptophysin-GFP and syntaxin-SBD-
GFP constructs, respectively. This work was funded by a Research Starter grant from the
PhRMA Foundation (SS), start-up funds from Case Western Reserve University (SS),
and the NIH Predoctoral Training Program in Molecular Therapeutics at Case Western
Reserve University (LB).
56
Chapter 3: Dynamic mechanisms of neuroligin-dependent presynaptic terminal
assembly in living cortical neurons
Luke A.D. Bury1
and Shasta L. Sabo1,2
Departments of 1Pharmacology and
2Neuroscience, Case Western Reserve University
School of Medicine, Cleveland, Ohio.
*Parts of this chapter were published in the journal Neural Development, Volume 9,
Issue 13, May 29, 2014.
57
Abstract
Synapse formation occurs when synaptogenic signals trigger coordinated
development of pre and postsynaptic structures. One of the best-characterized
synaptogenic signals is trans-synaptic adhesion. However, it remains unclear how
synaptic proteins are recruited to sites of adhesion. In particular, it is unknown whether
synaptogenic signals attract synaptic vesicle (SV) and active zone (AZ) proteins to
nascent synapses or instead predominantly function to create sites that are capable of
forming synapses. It is also unclear how labile synaptic proteins are at developing
synapses after their initial recruitment. To address these issues, we used long-term, live
confocal imaging of presynaptic terminal formation in cultured cortical neurons after
contact with the synaptogenic postsynaptic adhesion proteins neuroligin-1 or SynCAM-1.
Surprisingly, we find that trans-synaptic adhesion does not attract SV or AZ proteins nor
alter their transport. In addition, although neurexin (the presynaptic partner of neuroligin)
typically accumulates over the entire region of contact between axons and neuroligin-1-
expressing cells, SV proteins selectively assemble at spots of enhanced neurexin
clustering. The arrival and maintenance of SV proteins at these sites is highly variable
over the course of minutes to hours, and this variability correlates with neurexin levels at
individual synapses. Together, our data support a model of synaptogenesis where
presynaptic proteins are trapped at specific axonal sites, where they are stabilized by
trans-synaptic adhesion signaling.
58
Introduction
Upon axon-dendrite contact, the extracellular domains of axonal and dendritic
adhesion proteins interact, leading to recruitment of synaptic proteins and subsequent
synapse formation (5, 136, 138). A variety of synaptogenic adhesion partners have been
discovered, including neurexin/neuroligin, SynCAM/SynCAM, and neurexin/LRRTM2,
among others (117, 128, 129, 143, 163-166, 176, 214-216). Of these, the most studied
trans-synaptic pair is postsynaptic neuroligin and its interaction with presynaptic
neurexin.
Although a number of studies have focused on understanding the functions and
mechanisms of trans-synaptic adhesion molecules in recent years, it remains unclear how
synaptogenic adhesion results in presynaptic protein recruitment and synaptogenesis
(124, 217). Importantly, it is not known whether trans-synaptic adhesion actively attracts
synaptic proteins to sites of signaling or acts primarily to stabilize proteins at nascent
terminals. In addition, it is unknown whether synaptic protein recruitment proceeds
continuously until the site is saturated, or if SV and AZ protein levels are modulated even
at the earliest stages of development, as they are at mature synapses (218-225). Finally, it
is unclear if trans-synaptic adhesion regulates synaptic protein levels at individual
synapses or primarily creates sites in the axon that are capable of synapse formation
while other cellular processes regulate levels of recruitment.
To answer these questions, we induced synaptogenesis via contact with
neuroligin-1 and SynCAM-1, then employed short and long-term time-lapse confocal
imaging of presynaptic protein recruitment to nascent sites of trans-synaptic signaling.
59
This paradigm allowed us to record recruitment from the initiation of trans-synaptic
adhesion onward. We found that SV protein recruitment to individual sites of trans-
synaptic adhesion fluctuated, as did protein recruitment at axon-dendrite synapses. These
changes in synaptic protein levels strongly correlated with the amount of neurexin-1β at
individual synapses. However, trans-synaptic adhesion did not attract SV or AZ proteins
to contacts or significantly alter synaptic protein transport. Unexpectedly, neurexin alone
was not the primary signal for recruitment since synaptic proteins were specifically
recruited to sites of enhanced neurexin-1β clustering, even though neurexin was present
throughout the contact region. Overall, our data support a model of synaptogenesis in
which presynaptic proteins are trapped at, but not actively attracted to, specific axonal
sites, where they are stabilized by trans-synaptic adhesion signaling.
60
Results
Synaptic protein recruitment fluctuates frequently and rapidly during assembly of
individual presynaptic terminals
Contact between an axon and dendrite is one of the first steps of synapse
formation; however, it is difficult to predict when and where axo-dendritic contact will
occur and whether synapses will form at these sites. To overcome this challenge, we
developed a strategy for long-term imaging of synaptic protein recruitment to nascent
sites of presynaptic terminal formation, initiated by contact with neuroligin-1-expressing
HEK293 cells as a proxy for post-synaptic dendrites. Specifically, 7-10DIV cortical
neurons were sparsely transfected with synaptophysin-GFP, to label synaptic vesicle
protein transport vesicles (STVs), which deliver SV proteins to developing presynaptic
terminals (47-49, 51-53, 58, 68-71, 81, 87, 114, 115, 117, 210, 226, 227).We then imaged
STVs in individual axons that contacted neuroligin-1-expressing HEK293 cells for up to
25h after contact (Figure 3.1A), collecting multiple “short sequences” of rapid imaging
(every 10s for 7.5min) separated by 1-2.5h. The “short sequences” granted the ability to
track rapid STV movements, observe recruitment of individual STVs, and distinguish
between STVs that were stably recruited and those that were en route through the contact
site while preserving neuron health. With this approach, it was possible to resolve a full
time-course of presynaptic protein recruitment at individual sites of trans-synaptic
signaling in individual axons.
To quantify STV recruitment within individual axons, the integrated densities of
all STVs were summed in each axonal region that made contact with a neuroligin-1-
61
expressing HEK293 cell and compared to regions without contact in the same axon.
Since integrated density corresponds to the sum of pixel intensities, STVs with higher
integrated densities were brighter and/or larger, indicating more SV protein. Enrichment
was calculated by determining the difference between fluorescence (integrated density
per length of axon) inside and outside the contact region, divided by the sum of the
fluorescence inside and outside. Overall, in regions that displayed recruitment (see
Methods), synaptophysin-GFP was increasingly enriched at sites of trans-synaptic
adhesion over the first 10h of imaging (Figure 3.1B; n = 12 contacts), similar to previous
observations using immunocytochemistry to examine populations of axons (115). No
enrichment was observed over the same time course when axons were contacted with
HEK293 cells expressing only HcRed, with no neuroligin, indicating that the observed
recruitment was a specific result of trans-synaptic adhesion (Figure 3.1C; n = 5 contacts).
Enrichment at neuroligin contacts was due to increased SV protein recruitment to
contacts without substantial changes in SV protein outside areas of contact (Figure 3.1D).
Within contact areas, SV protein levels started to rise 2h after contact was initiated. This
“rising/recruitment phase” progressed until reaching a plateau 10h after contact initiation
(Figure 3.1D). However, recruitment at individual contacts was quite labile (Figure 3.1E).
Throughout the imaging period, absolute levels of recruited synaptophysin-GFP
increased or decreased from one hour to the next, even for axons that displayed
enrichment for most or all of the 24h imaging period. These substantial fluctuations
occurred during both rising and plateau phases. Importantly, variability in fluorescence
was not caused by focal drift, since we employed Perfect Focus correction to maintain the
focus (see Methods). Therefore, in individual axons, recruitment of proteins does not
62
consistently increase: rather, levels of recruited SV proteins fluctuate while remaining
elevated compared to neighboring axonal areas without trans-synaptic signaling.
63
Figure 3.1: Levels of synaptic vesicle protein enrichment at individual trans-
synaptic adhesion sites are modulated throughout recruitment. (A) Images from live,
long-term, time-lapse confocal imaging of an axon expressing synaptophysin-GFP
(green) and contacting an HEK293 cell (magenta and white outline) that expresses
neuroligin-1 + HcRed. The top panel shows an axon and HEK293 cell at 1h after contact.
In this panel, synaptophysin-GFP at the contact site appears white in the overlay. The
remaining panels show only synaptophysin fluorescence for clarity. Images were
collected 1h, 12.5h and 20h after contact was induced (scale bar = 5m). (B) Time course
of STV enrichment in axonal regions that contact neuroligin-1-expressing cells.
Enrichment corresponds to the difference between the total STV integrated density within
the contact region and outside of the contact, normalized to the total STV integrated
density throughout the axon and expressed as a percentage. Positive values indicate
enrichment inside the contact area. Individual points (cyan) represent mean values from
45 images collected at 10s intervals, beginning at the time indicated on the x-axis.
Dashed line, overall mean at each time point (n = 12 contacts). Overall, enrichment
64
increased gradually over the first 10h then remained elevated. Error bars = S.E.M. (C)
Over the same time course, STVs were not enriched at sites of contact between axons and
HEK293 cells expressing HcRed but no neuroligin. (D) Integrated density of
synaptophysin-GFP inside (solid line) and outside (dashed line) of contacts over time.
Red bar, rising phase of enrichment (2-7.5h after contact). Synaptophysin increased at
contacts while remaining stable outside contacts. (E) Enrichment for two axons. At
individual contacts, enrichment was highly dynamic throughout the imaging period.
Therefore, although STV enrichment increases at trans-synaptic adhesion sites overall,
enrichment levels for individual axons vary throughout the first 24h of development.
65
Fluctuations in recruited SV protein levels could occur through at least two
mechanisms (Figure 3.2A). Once formed, sites of STV recruitment could be stable while
the level of protein at each stable site fluctuates. Alternatively, the sites themselves could
form and disappear. These two mechanisms are not mutually exclusive, and our live
imaging revealed both types of variability. Synaptophysin-GFP levels fluctuated at
individual sites that were stable over several hours (Figure 3.2B). Fluctuations often
occurred very rapidly, within minutes. In addition, sites of stable recruitment often
persisted for multiple hours before completely, and sometimes immediately, disbanding
(Figure 3.2C). Instant formation of stable sites was also observed (Figure 3.2C).
To assess the prevalence of fluctuations in synaptophysin recruitment levels at
individual sites, we determined the number of sites of stable STV accumulation that
displayed fluctuations in synaptophysin-GFP levels over time. Stable accumulation sites
were defined as sites in the axon where synaptophysin-GFP signal was consistently
present over the course of at least one short imaging sequence. At 24.9% of individual
stable accumulation sites (n = 115), there were observable fluctuations of synaptophysin-
GFP within short imaging sequences. For stable accumulation sites that persisted over
multiple short imaging sequences, synaptophysin-GFP levels changed between short
imaging sequences 18.4% of the time (n = 103). It should be noted that these are most
likely conservative estimates of the actual synaptic vesicle protein level fluctuations at
stable sites, as small fluctuations of protein at these sites are difficult to observe.
Additions and losses of entire stable sites of recruitment resulted in net changes in
the density of discrete, stable recruitment sites from one short sequence to the next (i.e.
over 1-2.5 hours) in 76.6% of imaging sequences during the rising phase and 73.8%
66
during the plateau phase. This indicates that sites of recruitment were highly labile
throughout synapse assembly. During the rising phase of recruitment, the additions and
losses resulted in net increases in the density of stable sites of recruitment in 54.7% of
imaging sequences (Figure 3.2D; n = 64). In contrast, during the plateau phase,
appearances and disappearances were more balanced, with net increases in discrete
contact sites only 38.1% of the time (Figure 3.2D).
67
68
Figure 3.2: Modulation of synaptic vesicle protein recruitment occurs through two
distinct mechanisms. (A) Model kymographs illustrating that variability in protein levels
could arise from fluctuations in synaptic protein levels at individual recruitment sites
(left) or through the addition/subtraction of entire recruitment sites (right). (B, C)
Kymographs of STVs (green) in axons that contact neuroligin-1-expressing HEK-293
cells (magenta). (B, box) Synaptophysin-GFP levels at stable contacts are variable over
the course of hours and even minutes (asterisks). (C, top box, top line) Sites of
recruitment that are stable over the course of hours can be eliminated, sometimes rapidly.
(C) To replace these sites, individual STVs can be captured (bottom line) and stabilized
(bottom box) at contact sites. t = time after contact, scale bars = 5m. (D) Histograms of
net changes in the densities of individual stable recruitment sites from one short imaging
sequence to the next. Stable sites were defined as puncta that appeared at a given site
throughout at least one short imaging sequence. The number of stable sites of recruitment
varied at the majority of contacts during both the rising and plateau phases, with a bias
toward puncta addition during the rising phase (left) but not the plateau phase (right).
69
Levels of synaptic vesicle proteins are highly labile at developing axo-dendritic
contacts
To determine if the same types of fluctuations in SV protein recruitment occur at
neuron-neuron synapses, we imaged regions of contact between axons expressing
synaptophysin-GFP and morphologically identified dendrites and somas expressing RFP.
Contacts were imaged for 12-24h at 7-10 DIV with the imaging protocol described above
(Figure 3.3A). Axo-dendritic and axo-somatic synapses exhibited many of the same
dynamics observed in axons contacting neuroligin-1-expressing HEK293 cells. Levels of
synaptophysin-GFP at individual sites of stable recruitment varied over time (Figure
3.3B,C). Synaptophysin-GFP protein levels at contact sites fluctuated 10.9% of the time
(n = 43) between short imaging sequences (over 1-2.5h) and 17.6% of the time (n = 49)
within short sequences (over 7.5min). Furthermore, individual recruitment sites were
established and dissolved over the course of hours (Figure 3.3B, C), with net changes in
the density of stable sites of recruitment occurring in 55.5% of imaging sequences
(Figure 3.3D; 31.1% additions and 24.4% losses; n = 45). These data indicate that the
dynamics of synaptic vesicle protein recruitment at axo-dendritic synapses and hemi-
synapses occurred over a similar time course. The similarities between recruitment
dynamics at spontaneous neuron-neuron synapses and neuroligin-1-induced sites are
especially striking since sites of recruitment may have been more mature in some neuron-
neuron synapses, and synapse formation between neurons may involve additional
signaling pathways. Together, the above data suggest that developing synapses undergo
extensive fluctuations in presynaptic protein levels, even during initial formation.
70
Figure 3.3: Synaptic vesicle protein recruitment to axo-dendritic contacts is similar
that at induced trans-synaptic signaling sites. (A) Image of an axon transfected with
synaptophysin-GFP (green) contacting somato-dendritic region from an adjacent cell
transfected with RFP (magenta). White box corresponds to contact region represented by
71
kymograph in (B). (B, C) Kymographs of STVs (green) in axons that contact RFP-
expressing dendrites from different neurons (red). Boxes, as seen in induced axo-
dendritic contacts, synaptophysin-GFP levels are variable at individual stable sites in
neuron-neuron contacts over the course of hours and minutes (asterisks). t = time after
imaging begins, scale bars (A) = 10m, (B, C) = 5m. (D) Histogram of net changes in
the densities of individual stable recruitment sites from one short imaging sequence to the
next for neuron-neuron contacts. The number of stable sites of recruitment fluctuated
between the majority of imaging sequences (corresponds to all non-zero bins), with a
small bias toward addition of stable sites of recruitment.
72
Synaptophysin is preferentially stabilized at clustered neurexin
Neuroligin induces synaptogenesis through its presynaptic binding partner
neurexin (5, 228). To ascertain the time course of neurexin clustering at contacts,
neuroligin-1-expressing HEK293 cells were added to neuronal cultures transfected with
neurexin-1β-GFP. Then contacts between neuroligin-1-expressing HEK293 cells and
axons expressing neurexin-1β-GFP were imaged immediately. Prior to contact, neurexin-
1β-GFP appeared dim and diffuse within the axon. Upon contact with neuroligin-1-
expressing HEK293 cells, neurexin-1β-GFP began to be recruited to sites of contact
within minutes, and showed strong recruitment within 1-2 hours (Figure 3.4A-C, n = 14
contacts), consistent with previous data (115). Because we have not found any neurexin-β
selective antibodies for immunofluorescence, we could not determine the extent to which
neurexin-1β-GFP was over-expressed in these experiments. However, similar
fluorescently tagged neurexin-1β-GFP has been used in a number of studies to investigate
the localization, trafficking and function of neurexin (229-232). Importantly, the
distribution pattern of α-neurexin is not significantly influenced by the level of neurexin
expressed (232). Therefore, although over-expression of neurexin in neurons increases
synapse density (230), it not expected to change the dynamics or distribution of neurexin.
Neuroligin-1 recruited neurexin-1β-GFP to the entire contact region, where
neurexin appeared in both punctate and diffuse forms, even within the same contacts
(Figure 3.4B). Recent work has shown that the synaptogenic effect of neuroligin depends
on the postsynaptic scaffolding protein S-SCAM, which works, in part, by clustering
neuroligin (216). To determine if post-synaptic clustering of neuroligin facilitates
presynaptic clustering of neurexin, HEK293 cells that co-expressed S-SCAM and
73
neuroligin-1 were dropped onto neurons that were transfected with neurexin-1β-GFP.
This led to enhanced clustering of neurexin-1β-GFP in axons that contacted transfected
HEK293 cells (Figure 3.4B), suggesting that the synaptogenic effects of S-SCAM-
induced neuroligin clustering might be due to enhanced clustering of presynaptic
neurexin. Indeed, artificial clustering of neurexin is sufficient to induce presynaptic
protein recruitment (124), and recent work suggests that neurexin clustering is critical for
the synaptogenic function of the neurexin-neuroligin interaction in Drosophila (125).
This raises the question of whether synaptic proteins are preferentially recruited to
punctate clusters of neurexin or whether they can be recruited to any site with neurexin
enrichment. To test this, neurons were co-transfected with neurexin-1β-GFP and
synaptophysin-RFP. Neuroligin-1-expressing HEK293 cells were then added to these
cultures, and contacts were imaged for up to 24h, as described above. Although there
was a combination of diffuse and punctate recruitment of neurexin-1β-GFP to contacts
(Figure 3.4A-C), stable sites of synaptophysin-RFP recruitment were almost exclusively
co-localized with neurexin-1β-GFP puncta (Figure 3.4C). Surprisingly, neurexin-1β-GFP
clustering and synaptophysin-RFP clustering occurred nearly simultaneously at most
shared sites (Figure 3.4D, n = 13/18 clusters). Neurexin clustered concurrently with SV
protein recruitment even when HEK293 cells expressed both neuroligin-1 and S-SCAM
to enhance neurexin clustering. These data support a model in which discrete neurexin
clustering is critical for stable, rapid, neuroligin-induced presynaptic protein recruitment
in mammalian cortical neurons.
74
Figure 3.4: Synaptic vesicle protein accumulation occurs at clustered neurexin and
correlates with levels of clustered neurexin. (A) Time-lapse images of an axon
transfected with neurexin-1β-GFP (green) contacting a neuroligin-1-expressingHEK293
cell (white outline). Neurexin is recruited to contact sites within minutes and enhanced at
these sites as imaging progresses. (B) Images of two axons that are transfected with
neurexin-1β-GFP (green) and contact cells expressing neuroligin-1 (left, white outline) or
neuroligin-1+S-SCAM (right, white outline). At neuroligin-1-only contacts, neurexin
appears both punctate (arrows) and diffuse (bars). Addition of S-SCAM enhanced
neurexin-1β-GFP clustering and reduced diffuse neurexin-1β-GFP labeling at contacts
75
(arrow). (C) Persistent sites of synaptophysin-RFP (magenta) recruitment appeared at
sites of punctate neurexin-1β-GFP (green) recruitment (arrows). t = time after contact
established. (D) Kymographs of neurexin-1β-GFP and synaptophysin-RFP, starting
immediately after contact with a neuroligin-1-expressing cell was established. Neurexin
and synaptophysin clustered simultaneously at the same sites in the axon (arrows). (E) At
an individual recruitment site, synaptophysin-RFP and neurexin-1β-GFP levels were
positively (r = 0.98) and significantly (p = 2.0x10-5
) correlated over several hours of
imaging. This correlation was significant for 11 of 14 co-recruitment sites that were
present for at least 5h. (F) When combining data from all recruitment sites in the same
axon, synaptophysin and neurexin integrated densities were also correlated (r = 0.91, p =
6.0x10-20
). Similar correlation was observed in 5 of 7 axons. (G) When the data from all
axons were pooled, neurexin and synaptophysin were positively and significantly
correlated (p = 1.0X10-74
, r = 0.82). (A-C) scale bars = 10m, (D) scale bar = 5m, (E-G)
correlations made from non-transformed data and plotted in log-log form for display
purposes.
76
Actin polymerization is occasionally co-localized with neurexin clustering
While the neuroligin-neurexin interaction by itself is capable of generating
synaptogenic signaling, other signaling pathways can interact with and enhance
neuroligin-neurexin-induced synapse formation. Specifically, n-cadherin can act in
concert with neuroligin to enhance synaptic vesicle recruitment to developing presynaptic
terminals (216, 233). N-cadherin can mediate its intracellular signaling through actin, and
actin is known to play a role in the recruitment of presynaptic components to the
developing synapse (234-236). Therefore, to determine whether actin plays a role in
mediating the initial recruitment of presynaptic components to synapses, the time-course
of actin polymerization to novel sites of neurexin signaling was determined. Specifically,
HEK293 cells expressing neuroligin were dropped onto 6-8 DIV neuronal cultures that
co-expressed neurexin-GFP and Lifeact-mcherry, which is a fluorescently-tagged small
peptide that binds endogenous F-actin (237, 238). HEK293 cells that contacted neurexin-
GFP and Lifeact-mcherry-expressing axons were then imaged immediately via live time-
lapse confocal microscopy at a frame rate of one frame per minute. Neurexin recruitment,
including neurexin clustering, was observed at multiple contact sites after 60 minutes.
Approximately half of the contact sites (n = 4/9 contacts in 4 separate axons) displayed F-
actin clustering 60 min. after the initiation of neurexin recruitment (data not shown).
The synaptic scaffolding molecule S-SCAM physically links n-cadherin with
neuroligin via intracellular interactions on the postsynaptic side of the synapse (216).
This linkage induces enhanced clustering of neurexin and enhanced synapse formation
(216)(Figure 3.4B). Since n-cadherin binding can mediate downstream actin
polymerization (235), we hypothesized that postsynaptically localizing n-cadherin with
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neuroligin might enhance actin polymerization presynaptically, particularly at sites of
neurexin clustering. Therefore, HEK293 cells expressing neuroligin + S-SCAM were
dropped onto 7-8 DIV axons that co-expressed neurexin-GFP and Lifeact-mcherry and
immediately imaged. Importantly, n-cadherin is endogenously expressed in HEK293
cells (239). Neurexin clustering was enhanced at these contact sites, compared to contacts
with HEK293 cells that did not express S-SCAM. However, co-localized actin
polymerization and neurexin clustering was observed in only 33% of contacts (data not
shown, n = 4/12 contacts in 3 separate axons).
While it is tempting to infer that actin is not typically involved in the initial
recruitment of synaptophysin, these experiments were performed in a limited number of
neurons. In previous experiments, simultaneous co-clustering of neurexin and
synaptophysin were not always observed (n = 5/18 contacts), suggesting that specific
neurons or even specific sites within the same neuron are more or potentially less capable
of this rapid co-clustering. Follow-up experiments involving additional neurons and
HEK293 cell/axon contacts are needed to confirm these results both with and without S-
SCAM as a postsynaptic organizer.
Levels of recruited synaptophysin and neurexin are strongly correlated at
developing presynaptic terminals
Our observation that SV proteins and neurexin selectively and simultaneously co-
cluster raises the question of whether synaptophysin recruitment and neurexin clustering
are coordinated. To test this, we first determined whether synaptophysin and neurexin
levels were correlated at individual sites of recruitment. Indeed, synaptophysin levels
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were positively and significantly correlated with neurexin levels at individual co-clusters
that were present for at least five consecutive short sequences of imaging (Figure 3.4E, p
< 0.05 at 11/14 sites from 7 axons, r = 0.37-0.98). Neurexin-1β-GFP and synaptophysin-
RFP levels were also significantly correlated when all clusters from individual axons
were pooled (Figure 3.4F, p < 0.05 in 5/7 axons, r =0.52-0.96) or when all axons were
combined (Figure 3.4G, n = 307 sites, p = 1.0X10-74
, r = 0.82). Therefore, recruitment of
SV proteins and clustered neurexin are coordinated during presynaptic terminal
assembly.
Next, we asked whether the long-term dynamics of synaptophysin and neurexin
recruitment were similar. To test this, 7-10 DIV neurons were transfected with either
synaptophysin-GFP and neurexin-tdTomato or the near-infrared fluorescent protein
RFP670 (Figure 3.5) (240). Contacts between axons that expressed both synaptophysin-
GFP and neurexin-tdTomato and dendrites that expressed RFP670 were then imaged over
the course of 12-25h. Like synaptophysin, levels of neurexin fluctuated significantly over
the course of long-term imaging at both spontaneously-formed axo-dendritic and axo-
somatic neuron-neuron contacts (Figure 3.5A,B) as well as contacts formed with
neuroligin-expressing HEK293 cells (Figure 3.5C). In addition, many neurexin-1β-GFP
clusters formed and disappeared over the course of hours (Figure 3.4C and 3.5),
resembling the formation and dissolution of stable sites of synaptophysin recruitment
(Figure 3.2 and 3.5).
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Figure 3.5: Neurexin and synaptophysin levels fluctuate over the course of minutes
at axo-dendritic contact sites and neuroligin induced developing synapses. (A) Image
of an axon co-expressing synaptophysin-GFP (green) and neurexin-tdTomato (red) and
contacting the somato-dendritic region of a neuron expressing RFP670 (blue). White box
corresponds to contact region represented by kymographs in (B). (B) Individual
kymographs of an axon expressing synaptophysin-GFP (top) and neurexin-tdTomato
(middle) that contacts a dendrite expressing RFP670 (bottom). Levels of both neurexin
and synaptophysin fluctuate over the course of minutes to hours at sites that contact
somato-dendritic regions of the adjacent neuron (arrows, top), including the formation
(white box) and disappearance (white arrowhead) of co-clusters. (C) Kymographs of an
axon expressing neurexin-GFP (left) and synaptophysin-RFP (right) contacting a
neuroligin-expressing HEK293 cell. The amount of neurexin and synaptophysin at co-
clustered sites (arrow, bottom) fluctuates extensively over hours and even within minutes
(white arrowheads). Scale bars (A,B) = 10m, scale bar (C) = 5m.
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Trans-synaptic signaling recruits synaptic proteins without substantially altering
their transport
It is not known how STVs disengage from their microtubule tracks and become
incorporated into synapses. Various signals that affect synaptogenesis also affect STV
transport (51), and it has been suggested that neuroligin-neurexin signaling might recruit
synaptic proteins by altering synaptic protein transport (241). Therefore, we next tested
the hypothesis that trans-synaptic signaling recruits presynaptic proteins by altering STV
transport. To test this hypothesis, synaptophysin-GFP was imaged every 10s for 7.5 min
at 1-4 hours after contact was established between axons and neuroligin-1-expressing
HEK293 cells (Figure 3.6A). Because STV transport is characterized by bursts of
movement interrupted by pauses (47, 48, 51, 53, 58, 87, 210), we measured how fast
STVs moved and how long they paused in the presence and absence of neuroligin-1. We
found that STV pause duration increased at contacts with neuroligin-1-expressing cells
when compared to contacts with HcRed-expressing control cells (Figure 3.6B; neuroligin
= 145.3s, +/- 20.8, n = 40 pauses; no neuroligin = 85.7s, +/- 15.5, p = 0.0217, n = 42
pauses). However, there was no neuroligin-1-dependent change in instantaneous velocity
(Figure 3.6B; neuroligin = 0.229m/s, +/- 0.023, n = 40 movements; no neuroligin =
0.196m/s, +/- 0.015, n = 56 movements, p = 0.391).
To investigate active zone protein transport, we looked at Piccolo-Bassoon
transport vesicles (PTVs), which deliver active zone proteins to developing presynaptic
terminals (49, 59-62, 73, 74, 81, 83, 87, 115, 118). STVs and PTVs can be transported
together in the axon (49, 87, 96). Therefore, neuroligin-neurexin interactions could alter
PTV transport, which in turn could recruit STVs. Similar to STVs, neuroligin-1 had no
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effect on the instantaneous velocity of PTVs (Figure 3.6C; neuroligin = 0.140m/s, +/-
0.008, n = 22 movements; no neuroligin = 0.158m/s, +/- 0.009, n = 59 movements, p =
0.473). Unlike STVs, contact with a neuroligin-1-expressing cell had no effect on the
pause duration of PTVs (Figure 3.6C; neuroligin = 173.1s, +/- 24.9s, n = 32 pauses; no
neuroligin = 145.5s, +/- 14.0s, n = 53 pauses, p = 0.634). Therefore, neuroligin-neurexin
signaling does not recruit STVs indirectly via regulation of PTV transport.
To determine if other forms of synaptogenic trans-synaptic signaling alter STV
movement, we also performed the live-imaging assay described above using HEK293
cells transfected with SynCAM-1, a trans-synaptic adhesion molecule that possesses
synaptogenic properties similar to neuroligin-1(129, 143, 156). Unlike neuroligin-
neurexin signaling, SynCAM-1-mediated trans-synaptic adhesion did not alter STV pause
duration (Figure 3.6D; SynCAM = 112.5s, +/- 14.0, n = 53 pauses; no SynCAM =
125.5s, +/- 13.4, n = 58 pauses, p = 0.332). However, SynCAM-1 signaling caused a
small increase in the instantaneous velocity of STVs (Figure 3.6D; SynCAM =
0.235m/s, +/- 0.021, n = 52 movements; no SynCAM = 0.183m/s, +/- 0.012, n = 58
movements, p = 0.030). Since the time-course of presynaptic protein recruitment to
SynCAM-1 adhesion sites has not been established, we also imaged STVs at 24h after
contact with SynCAM-1. After 24h, SynCAM-1-expressing cells had no effect on STV
movement (Figure 3.6E; pause duration: SynCAM = 140.3s, +/- 15.6, n = 62 pauses; no
SynCAM = 116.5s, +/- 14.4, p = 0.197, n = 71 pauses, instantaneous velocity: SynCAM
= 0.202m/s, +/- 0.012, n = 71 movements; no SynCAM = 0.171m/s, +/- 0.007, p =
0.060, n = 86 movements).
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Neuroligin and SynCAM interact with different presynaptic adhesion molecules
(5, 136, 242). Therefore, it might be expected that the effects of neuroligin and SynCAM
would be additive or synergistic. However, contact with HEK293 cells that expressed
both neuroligin-1 and SynCAM-1 had no effect on the movement of STVs (Figure 3.6F;
pause duration: neuroligin/SynCAM = 124.6s, +/- 12.7, n = 61 pauses; no
neuroligin/SynCAM = 116.7s, +/- 14.3, p = 0.482, n = 51 pauses; instantaneous velocity:
neuroligin/SynCAM = 0.196m/s, +/- 0.012, n = 63 movements; no neuroligin/SynCAM
= 0.175m/s, +/- 0.008, p = 0.187, n = 73 movements).
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Figure 3.6: Trans-synaptic signaling has little effect on synaptic vesicle or active
zone protein transport. (A) Time-lapse images of a neuroligin-1 + HcRed-expressing
HEK293 cell (magenta) contacting an axon expressing synaptophysin-GFP (green).
Arrows, position of individual STV in each frame. Scale bar = 10m. (B) Contact with
neuroligin-1-expressing cells increases the mean pause duration but not instantaneous
velocity of STVs inside the contact region. (C) For PTVs, neither the mean pause
duration nor the instantaneous velocity was affected by neuroligin-1-induced trans-
synaptic signaling. (D) Contact with SynCAM-1-expressing cells for 1-4h did not affect
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STV pause duration but increased their instantaneous velocity. (E) Contact with
SynCAM-1 for 24h had no effect on STV pause duration or instantaneous velocity. (F)
Contact with both neuroligin-1 and SynCAM-1 had no effect on STV pause duration or
instantaneous velocity. *, p < 0.05; t = time after imaging begins; error bars = S.E.M.
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In addition to comparing the transport of STVs and PTVs in axonal regions that
contact HEK293 that either did or did not express synaptogenic adhesion molecules, we
also compared axonal transport inside and outside of contact regions within the same
axon. Each combination of transport vesicle and non-neuronal cell treatment described
above was tested. No significant (all p > 0.05) difference in average pause duration or
instantaneous velocity was observed when comparing transport vesicles inside of contact
regions vs. those outside of contact regions (data not shown), with one exception. The
average pause duration of STVs was significantly increased inside axonal regions that
contacted SynCAM-expressing cells for 24h (outside contact = 105.6s +/- 6.1, n = 339
pauses; inside contact = 140.3s +/- 15.6, n = 62 pauses).
Together, our data suggest that while trans-synaptic signaling might have some
effect on STV movement, regulation of transport is not the main mechanism through
which trans-synaptic adhesion mediates synaptic protein recruitment. It is unclear
whether the distinct effects of neuroligin and SynCAM on STV trafficking represent
mechanistic differences in how recruitment of STVs occurs down-stream of presynaptic
neurexin and SynCAM. Since neuroligin plus SynCAM did not result in the same
changes as either neuroligin or SynCAM, it is possible that either (i) the observed
changes are not essential for STV recruitment, or (ii) when accumulated at the same sites,
presynaptic neurexin and SynCAM compete with each other for some factor that is
necessary for the observed effects on STV trafficking, essentially dampening their
distinct effects.
Synaptic vesicle and active zone protein transport vesicles are not attracted to sites
of trans-synaptic signaling
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Synaptogenic adhesion could recruit presynaptic machinery by actively attracting
synaptic proteins to sites of trans-synaptic signaling. To test this hypothesis, we
determined whether STV movements are biased towards sites of neuroligin-1 contact
(Figure 3.7). First, we quantified the net distance and direction moved for individual
STVs. Movements toward the contact were assigned positive values, while movements
away from the contact were negative. This analysis indicated that neuroligin-1 signaling
did not bias net STV movement toward neuroligin-1 (Figure 3.7B; net movement = -
0.078 +/- 0.857m, n = 67 STVs), similar to contacts with HcRed-expressing HEK293
cells (Figure 3.7C; net movement = -1.39 +/- 1.22m, n = 44 STVs; p = 0.60). Since
trans-synaptic signaling might only attract vesicles near the contact, we then restricted the
analysis to STVs with initial positions adjacent to the contact, within a region equal to the
contact in length. Again, neuroligin-neurexin signaling did not bias STV transport
(Figure 3.7D; Nlgn = 0.000 +/-0.888m, n = 17 STVs; HcRed = -3.08 +/- 1.69m, n = 19
STVs; p = 0.23). Moreover, the distributions of the net movements were not altered by
neuroligin-neurexin signaling (p > 0.15, Kolmogorov-Smirnov). In addition, neuroligin-1
did not change the fraction of STVs moving toward contacts (Figure 3.7E; # STVs
moving toward/# STVs moving away: Nlgn = 0.89, n = 66 STVs; HcRed = 0.83, n = 44
STVs). Similarly, STVs adjacent to contacts were not attracted to contacts (Figure 3.7F;
Nlgn = 0.89, n = 17 STVs; HcRed = 0.46, n = 19 STVs).
STVs also displayed no bias in movement towards HEK293 cells expressing
SynCAM-1- or SynCAM-1+neuroligin-1. SynCAM-1did not alter the net distance and
direction moved by all imaged STVs (p = 0.33, n = 136 and 88 STVs for SynCAM and
HcRed, respectively) or STVs adjacent to contacts (p = 0.23, n = 39 and 28 STVs for
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SynCAM and HcRed). The same was true when contacts expressed both SynCAM-1 and
neuroligin-1 (all STVs: p =0.70, n = 136 and 69 STVs for SynCAM+neuroligin and
HcRed, respectively; near contacts: p = 0.55, n = 39 and 18 STVs for
SynCAM+neuroligin and HcRed, respectively). Likewise, STV movement was not
biased toward contacts with SynCAM-1after 24h of contact (p = 0.13, n = 39 and 36
STVs for SynCAM and HcRed, respectively). Again, the data were not significantly
different when the distributions were compared using the Kolmogorov-Smirnov test (p >
0.11). Furthermore, the ratio of STVs moving toward vs. away from contact regions (for
SynCAM, SynCAM+neuroligin, or 24hr SynCAM) was less than or equal to 1.13,
indicating a lack of attraction to sites of trans-synaptic adhesion. Taken together, the
above data demonstrate that trans-synaptic signaling does not actively recruit STVs from
outside regions of contact.
Although STVs were not actively attracted to contact sites, it remained possible
that trans-synaptic signaling actively recruits active zone precursors to sites of contact,
followed by passive capturing of STVs. To test this, we determined whether movements
of PTVs were biased toward trans-synaptic adhesion sites. Similar to STVs, PTVs
showed no bias in movement towards neuroligin-1 contact sites (Figure 3.7G), both when
including all PTVs in the analysis (Nlgn = -0.35+/-0.56m, n = 79 PTVs; HcRed =
1.38+/-0.68, n = 69 PTVs; p = 0.071) and when limiting the analysis to PTVs with initial
positions adjacent to contacts (Nlgn = -1.34+/-1.12m, n = 26 PTVs; HcRed = 0.72+/-
0.86, n = 28 PTVs; p = 0.046). When the distributions were compared using the
Kolmogorov-Smirnov test, no differences were seen (p > 0.084). Collectively, our data
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indicate that trans-synaptic adhesion signaling does not attract synaptic vesicle or active
zone matrix proteins to developing synapses.
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Figure 3.7: Trans-synaptic signaling does not attract synaptic vesicle or active zone
proteins. (A) Schematics of HEK293 cell contact with an axon. Analysis was done on all
STVs with initial positions either outside the contact area (gray area, left, center) or
directly adjacent to the contact region (gray area, right). (B-D) Histograms of the net
movement of STVs towards (positive) or away from (negative) contacts with HEK293
cells expressing either neuroligin-1 + HcRed (B, D) or HcRed only (C). STV movement
was not biased towards sites of neurexin-neuroligin signaling, for either all STVs or only
STVs adjacent to the contact. (E, F) Scatter plot of the number of STVs moving towards
vs. away from contacts with cells expressing neuroligin-1 + HcRed (blue circles) or
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HcRed only (red squares). Each small point represents the ratio for one axon, with larger
points representing additional axons with the identical ratio. These ratios were
approximately the same for axons that contact cells expressing neuroligin-1 or HcRed
(black line = unity ratio line). (G) Similar to STVs, PTVs were not attracted to sites of
neuroligin-1 contact. (B-D, G) Red arrows = mean.
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Initial arrival of STVs and PTVs to a developing synapse is likely uncoordinated
Due to the variety of synaptic components transported by STVs and PTVs, it is
possible one type of transport vesicle is initially recruited to a developing synapse and
subsequently traps the other type of transport vesicle and/or additional transport vesicles
of the same type. If this is the case, then one of these transport vesicles would always
arrive at a developing synapse before the other. Therefore, to determine whether STVs or
PTVs are recruited to novel presynaptic terminals first, 6-8 DIV neurons were transfected
with either RFP670 or bassoon-GFP + synaptophysin-RFP. Axons that co-expressed
bassoon-GFP and synaptophysin-RFP and made contact with RFP670-expressing
dendrites were then imaged at a rate of 1 frame per 90s for up to 24 hours. During this
time, newly stabilized presynaptic terminals were formed along the axo-dendritic contact.
Synapse stabilization was defined as the co-recruitment of synaptophysin-RFP and
bassoon-GFP to the same site in the axon where both components remained at that site
continuously for at least one hour.
Overall, 51 newly stabilized presynaptic terminals were observed at 11 unique
neuron-neuron contacts. Upon visual examination of the images, it appeared that
synaptophysin-GFP arrived at the synapse first 31% of the time (n = 16/51 stabilized
synapses), bassoon-GFP arrived at the synapse first 33% of the time (n =17/51 stabilized
synapses), and simultaneous recruitment of synaptophysin-GFP and bassoon-GFP
occurred 35% of the time (n = 18/51 stabilized synapses, data not shown). This implies
that there is no defined order for when STVs or PTVs arrive at the synapse. In addition,
the high proportion of simultaneous arrivals suggests that when synaptogenic signaling
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occurs, both STVs and PTVs can be recruited to the same site rapidly, at least within 90
seconds of each other, as that was the length of time between imaging frames.
There are caveats to this analysis, however. The frame rate of 1 frame per 90s was
utilized to maximize temporal resolution while maintaining cell health over long periods
of time. Long-duration imaging was necessary because novel formation of presynaptic
terminals is relatively rare on existing axo-dendritic contacts. It is possible that one type
of transport vesicle is always stabilized first and then rapidly recruits the other type of
vesicle into the synapse. In addition, although there were many instances of STVs or
PTVs arriving to a new synaptic site first, it was often difficult to distinguish when one of
these components was initially stabilized at a specific site because both STVs and PTVs
appeared to pause at these sites before becoming more “permanently” fixed to the site.
This poses a challenge for analysis, as it would be impossible to distinguish between a
transport vesicle that pauses at the same site every 90s during the imaging of each frame
and a truly stabilized synaptic component. Therefore, higher temporal resolution is
needed to distinguish between these two phenomenon. Ongoing experiments will utilize
the synapse induction and long-term imaging protocol described in Figures 3.1, 3.2, and
3.3 to address this issue.
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Materials and Methods
All studies were conducted with an approved protocol from the Case Western
Reserve University Institutional Animal Care and Use Committee, in compliance with the
National Institutes of Health guidelines for the care and use of experimental animals.
Neuronal culture and transfection
Primary neuronal cultures were prepared from rat or mouse cortices on 18mm
glass coverslips as described previously (50, 51, 87, 207) and maintained in Neurobasal
A media with B27 Supplement (Invitrogen, Carlsbad, CA, USA). At 6-9 days in vitro
(DIV) and 24-48h prior to imaging, neurons were transfected using Lipofectamine 2000,
essentially according to the manufacturer’s instructions (Invitrogen). DNA was used at
1μg per 18mm coverslip with the exception of GFP-bassoon, which was transfected at
2μg per 18mm coverslip due to its large size. This amount of DNA results in efficient
labeling of transport vesicles without significant over-expression within axons (50, 51,
53, 87). Even so, neurons were chosen for imaging that displayed moderate expression
levels of the transfected constructs to avoid any possible effects of over-expression. For
double transfections, the localization and movement of fluorescently-tagged proteins
appeared similar to single transfections. In addition, synaptic vesicle protein transport
vesicles (STVs) labeled with synaptophysin-RFP and synaptophysin-GFP were similar in
size, movement, and localization.
Synaptophysin-GFP, GFP-bassoon (GFP-Bsn 95-3938), synaptophysin-mRFP,
synaptophysin-mcherry, and neurexin-1β-GFP were generous gifts of Drs. Jane Sullivan
(University of Washington, Seattle), Thomas Dresbach (Georg August University,
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Gottingen), Jurgen Klingauf (University of Muenster), Matthijs Verhage (Vrije
Universiteit, Amsterdam), and Camin Dean (The European Neuroscience Institute,
Gottingen). These constructs have been shown to be functional and properly localized
and have been used previously for studies of protein trafficking and localization (50, 61,
115, 211, 213, 243). Fluorescent proteins attached to bassoon and synaptophysin
effectively label PTVs/active zones and STVs/synaptic vesicles, respectively (48, 50, 53,
60, 61, 87, 115, 210, 211). RFP670 was purchased from Addgene [plasmid # 45457].
Neurexin-tdTomato was generated by seamlessly replacing the GFP sequence in the
neurexin-GFP construct with tdTomato via Gibson Assembly (New England
Biosystems).
HEK293 cell culture and transfection
HEK293 cells were cultured and maintained in DMEM supplemented with 10%
fetal calf serum and penicillin/streptomycin (Invitrogen, HyClone). Cells were
transfected 24-48h prior to dissociation and dropping onto neuronal cultures. A 3:2 or 2:1
ratio of trans-synaptic adhesion DNA to HcRed DNA (Clontech) was used to ensure co-
expression of both types of DNA constructs in the same HEK293 cell (51). To confirm
co-expression, HEK293 cells were transfected with SynCAM-GFP, HA-neuroligin-1, and
HcRed. After 48h, cells were fixed and labeled with anti-HA antibody to detect HA-
neuroligin-1. Over 85% of HEK293 cells transfected with HcRed also expressed
SynCAM-GFP and HA-neuroligin-1 (data not shown). HA-neuroligin-1 and HA-
SynCAM-1 were generous gifts of Drs. Peter Scheiffele (University of Basel) and Philip
Washbourne (University of Oregon, Eugene) and have previously been shown to recruit
synaptic proteins in the hemi-synapse assay (128, 244). Myc-tagged S-SCAM was
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provided by the labs of Yutaka Hata (Tokyo Medical and Dental University, Tokyo,
Japan) and Yoshimi Takai (Kobe University, Kobe, Japan) (245) and purchased from
Addgene (plasmid # 40213245). pDEST/Lifeact-mcherry-N1 was provided by Dr. Robin
Shaw (Cedars-Sinai, Los Angeles, CA, USA) (238) and purchased from Addgene
[plasmid # 40908].
Imaging
Imaging was performed with a C1 Plus confocal system on a Nikon Eclipse Ti-E
microscope utilizing a 40x Nikon Plan Apo 0.95NA objective. Lasers used for excitation
were 488nm argon and 543nm and 633 nm helium-neon, while detection filters were
515/30nm bandpass for GFP and 590/50nm bandpass for HcRed/mcherry/mRFP. For
multi-color imaging, channels were imaged sequentially to avoid bleed-through. To avoid
focal plane drift, the Perfect Focus System (Nikon) was used.
Long-term imaging
Transfected HEK293 cells were dissociated with Hank’s-based, enzyme-free cell
dissociation buffer (Invitrogen). Cells were resuspended in neuronal media then 8 X 104
cells were dropped onto each coverslip of transfected neurons. Contacts between
transfected axons and transfected HEK293 cells were imaged, with axons identified by
their morphological characteristics (50, 51). Dual-color images were collected every 10s
for 7.5min, with scan times for both frames not exceeding 3.3s. This interval yields high
temporal resolution of STV movement with minimal phototoxicity (87). This imaging
protocol was repeated every hour for the next 4h then every 2.5h for up to 25h. To ensure
that cells remained healthy, imaging was performed in a custom made environmental
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chamber held at 32°C. The pH was maintained at physiological levels by equilibration in
5% CO2 for 15-30min prior to sealing with vacuum grease (Dow Corning).
Short-term imaging
Synaptogenesis was induced as described above. After 1-4h, contacts between
transfected axons and transfected HEK293 cells were imaged every 10s for 7.5min with
constant perfusion of artificial cerebrospinal fluid (ACSF: 120mM NaCl, 3mM KCl,
2mM CaCl2, 30mM D-glucose, 20mM HEPES, and 0.2% sorbitol, pH 7.3) at 25°C. At
this temperature, STV movement is not significantly altered compared to STV movement
at physiological temperatures (51). In previous studies, the fastest velocities recorded for
STV movements were approximately 1μm/s (51, 53, 210), while maximal PTV velocities
were lower (60, 73). Vesicles moving at this maximal velocity would move 10μm
between frames during of imaging, a small fraction of the total axonal area typically
imaged. Therefore, the rapid imaging of STVs/PTVs performed here enabled reliable
identification and tracking of vesicles.
Neurexin-GFP recruitment
Axons expressing neurexin-GFP were imaged for at least 5min before contact
with HEK293 cells to identify neurexin-GFP that was previously clustered through
endogenous mechanisms. HEK293 cells transfected with HA-neuroligin-1 plus HcRed
were added to the imaged coverslip until at least one transfected HEK293 cell made
contact with the neurexin-expressing axons. This contact was then imaged for up to 2h at
a rate of 1 frame/min.
Analysis
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STVs and PTVs were manually tracked using custom written macros for ImageJ
(NIH, Bethesda, MD, USA), while blind to the locations of the HEK293 cells. Analysis
was restricted to axons that did not move during imaging and where at least one vesicle
moved within the imaged area. To facilitate tracking, imaging was conducted on axons
with an intermediate to low density of STVs or PTVs. Movements were quantified by
importing the positional data into Matlab and subjecting them to custom written analysis
(Mathworks, Natick, MA, USA). Pausing was defined as a period of at least 10s where
the velocity was lower than 0.1μm/s, as previously described (51, 87). Vesicles that did
not move during imaging were not included in this analysis, and analysis was restricted to
vesicles that could be tracked throughout the imaging period. Therefore, very long
pauses and vesicles that moved at high velocities for long periods of time might be
underrepresented. Considering the scan speed, pixel density, and average size of
transport vesicles, a typical STV or PTV was imaged within 10-50ms.
Integrated density analysis was performed using custom-written macros in ImageJ
that determined the location and integrated density of each transport vesicle on the axon
for each image in a sequence. Contact regions were defined as axonal regions that
overlapped with fluorescence from a transfected HEK293 cell. ‘% Enrichment’ was
determined by the following equation:
% Enrichment = [(IntDenin - IntDenout)/ (IntDenin + IntDenout)]*100
Where IntDenin corresponded to the mean integrated density per m of axon length inside
the contact, and IntDenout corresponded to the mean integrated density per m outside the
contact for the same axon. Contacts were considered to display recruitment when the %
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enrichment was (i) positive for at least 70% of time points 4 hours after contact was
initiated and (ii) greater than 10% for at least half of all time points 4+ hours after contact
initiation. Data are shown as the mean +/- standard error when applicable. Significance
was determined via Wilcoxon rank sum test unless otherwise indicated. For correlation
data, r = Pearson’s linear correlation coefficient and p-values were determined using
Student’s t-test.
Acknowledgements
We thank Michael Sceniak for helpful discussions and suggestions on the project. We
also thank Valentina Savchenko, Teresa Evans, Ruth Siegel, Lynn Landmesser, and
Brian McDermott for helpful reading of the manuscript. In addition, we thank Louie
Zhou and Jeesoo Kim for technical assistance, Ryan Yavorsky for assistance with
analysis and Corbett Berry for assistance with ImageJ programming. This work was
funded by National Institute of Mental Health Grant R01MH096908 (SLS), Simons
Foundation Autism Research Initiative (SLS), PhRMA Foundation Research Starter
Award (SLS) and the NIH Predoctoral Training Program in Molecular Therapeutics at
Case Western Reserve University (LADB).
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Chapter 4: Imaging presynaptic dynamics during mouse cortical development via
two-photon confocal microscopy
Teresa A. Evans1,2*
, Luke A.D. Bury1*
and Shasta L. Sabo1,2
Departments of 1Pharmacology and
2Neuroscience, Case Western Reserve University
School of Medicine, Cleveland, Ohio.
*For individual contributions, please see Materials and Methods
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Introduction
The previous experiments described in Chapters 2 and 3 have determined how the
transport of presynaptic components is coordinated prior to synapse formation and how
recruitment of these components progresses in response to induced synaptogenic
signaling. These experiments were done exclusively in living cultured cortical neurons,
which allowed for easy manipulation and high spatial and temporal resolution when
performing live, time-lapse confocal imaging. While this in vitro system offers many
benefits, it lacks key characteristics that are present in the intact brain. Specifically,
normal connectivity that develops between specific brain regions is not recapitulated in
neuronal cultures. In addition, many developing cortical circuits often require
environmental stimuli to refine their connectivity, which is also lacking in culture.
Understanding how the dynamic processes described above proceed within the
intact mammalian brain would therefore yield critical insight into how cortical circuits
are shaped. To achieve this insight, Cre-induced synaptophysin-tdtomato was
exogenously expressed in the mouse brain by viral injection of Cre-GFP. Cranial
windows were applied to these mice and synaptophysin-tdtomato was imaged within the
brain via 2-photon microscopy over the course of minutes. To detect whether presynaptic
dynamics change over the course of development, multiple imaging time-courses were
taken from the same animals on separate dates ranging from postnatal day 13 (P13) to
P22. These experiments are the first to detect fluorescently-tagged presynaptic proteins
within axons of the intact mammalian brain and add insight into synaptic dynamics that
occur during development.
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Results
To analyze presynaptic dynamics during development in vivo, presynaptic
terminals must be both labeled and detected within the brain. In order to fluorescently
label presynaptic terminals, “stop-flox” synaptophysin-tdtomato transgenic mice were
obtained. These mice possess an allele that contains a loxP-flanked stop codon that is
upstream of a synaptophysin-tdtomato fusion gene (Figure 4.1A). This stop codon
normally inhibits synaptophysin-tdtomato expression. However, with Cre expression, the
stop codon is removed and synaptophysin-tdtomato is expressed (Figure 4.1A). To
induce synaptophysin-tdtomato expression in the brain, an adeno-associated virus (AAV)
encoding Cre-GFP was injected via Hamilton syringe into the right hemisphere of
neonatal pups aged P2-P4. This limited expression to cell bodies located in the right
hemisphere of the cortex (Figure 4.1B). In addition, the expression of Cre-GFP was
driven by the synapsin promoter, which restricted expression to neurons.
In order to detect Cre-induced synaptophysin-tdtomato expression, cranial
windows were applied at postnatal day 10 to mice which had previously been injected
with Cre-GFP AAV. Windows were applied over the left hemisphere of the cortex in
order to image synaptophysin-tdtomato expression on the contralateral side of the
injection (Figure 4.1C). This restricted the observable synaptophysin-tdtomato signal to
callosally projecting axons in the left hemisphere whose somas were located in the right
hemisphere. This imaging strategy provided a number of distinct advantages compared to
imaging the ipsilateral hemisphere. First, since only axons cross the midline and project
to the contralateral side of the brain, the synaptophysin-tdtomato puncta that were
observed were strictly axonal and not mislocalized to dendrites. In addition, only
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excitatory neurons produce callosal projections, thereby limiting the synapses that are
detected to a specific population of cortical neurons. This neuronal population is
clinically relevant as well, as defects in callosal development have been linked to a
number of neurological disorders (246, 247).
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Figure 4.1: Synaptophysin-tdtomato expression in transgenic knock-in mouse. (A)
Schematic for synaptophysin-tdtomato expression in stop-flox synaptophysin-tdtomato
mice. Injection of AAV encoding Cre-GFP downstream of the synapsin promoter induces
Cre expression. In stop-flox synaptophysin tdtomato mice, this Cre expression drives
recombination of the loxP sites, thereby removing the stop codon and inducing
synaptophysin-tdtomato expression in neurons. (B) Image of brain expressing Cre-GFP
(green) in the right hemisphere (white circle). Cranial window application and
subsequent imaging occurs on the contralateral side (white square, arrow). (C) Example
of single image taken in living mouse via two-photon confocal imaging of
synaptophysin-tdtomato (yellow) through a cranial window. Scale bar = 20m.
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To determine if synaptophysin-tdtomato effectively localized to presynaptic
terminals in vivo, stop-flox synaptophysin-tdtomato mice that were injected with Cre-
GFP were perfused, sectioned, and labeled with antibodies against the presynaptic protein
synapsin. As highlighted by the white circles in Figure 4.2A, synaptophysin-tdtomato
colocalized frequently with synapsin labeling. To further confirm the synaptic
localization of synaptophysin-tdtomato, stop-flox synaptophysin-tdtomato mice were
cross-bred with Thy-1-YFP mice which express YFP in a sparse population of layer V
pyramidal cells. After injection with the Cre-GFP virus, the brains of these mice were
perfused and sectioned. As shown in Figure 4.2B, synaptophysin-tdtomato puncta
frequently and closely apposed the apical dendrite of YFP+ cells, suggesting a synaptic
localization. It should be noted that, although these data suggest a synaptic localization
for many, if not most synaptophysin-tdtomato puncta, it does not rule out the possibility
of some synaptophysin-tdtomato signal being non-synaptic and/or part of transport
vesicles within the axon.
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Figure 4.2: Synaptophysin-tdtomato is localized to presynaptic terminals. (A)
Synaptophysin-tdtomato (magenta) in the cortex co-localizes with antibody-labeled
synapsin (green). (B, left image) Multiple synaptophysin-tdtomato puncta (magenta) are
seen closely apposed to the apical dendrite of a YFP-filled cell (green). White circles and
square highlight co-localization points. (B, right images) Z-projections of the area
highlighted by the white square further indicate that synaptophysin-tdtomato and
dendrites are closely apposed in three-dimensional space. Scale bar for left image in (B)
= 10m, scale bar for right images in (B) = 1m.
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In order to monitor presynaptic dynamics within the intact brain, synaptophysin-
tdtomato was imaged through cranial windows in living mice via two-photon time-lapse
imaging. The frame rate of all time-lapse movies was one frame per thirty seconds, and
images were collected for up to one hour. In between each imaging frame, 30 seconds of
rest without imaging was performed to prevent overheating of the brain tissue. In some
instances, multiple time-lapse images of the same animal over the course of days were
obtained. This was not possible in all cases, however, as bone growth under the cranial
window occasionally obscured synaptophysin-tdtomato detection. Upon imaging, it was
determined that the majority of synaptophysin-tdtomato was detected in superficial areas
near the pial membrane, likely in layer I of the cortex. Therefore, our analysis focused on
these synapses which were consistently and strongly labeled across multiple animals over
multiple days of imaging. In order to determine whether the dynamics of presynaptic
terminals in this cortical layer change during development, imaging was done in animals
ranging in age from P13-P22, a time period in which critical synaptic development takes
place (248, 249). Movies were then binned according to specific developmental age.
Both splitting and merging of individual synaptophysin-tdtomato puncta were
observed at all points in development (Figure 4.3A,B). Both the percent of puncta that
split per minute of imaging and the percent of puncta that merged per minute of imaging
increased during the P16-P19 time period (compared to P13-P15) before becoming
reduced back to near original levels during the P20-P22 time period (for all percentages,
value = (# split puncta in time-lapse series/ total puncta in time-lapse series / length of
time-lapse series): for P13-P15, % puncta split per minute = 0.038, +/- 0.013, n = 6 time-
lapse series; % puncta merged per minute = 0.067, +/- 0.027, n = 6 time-lapse series; for
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P16-P19, % puncta split per minute = 0.14, +/- 0.032, n = 14 time-lapse series; % puncta
merged per minute = 0.18, +/- 0.034, n = 14 time-lapse series; for P20-P22, % puncta
split per minute = 0.066, +/- 0.018, n = 5 time-lapse series; % puncta merged per minute
= 0.11, +/- 0.042, n = 5 time-lapse series). The rates for both processes were quite similar
at each time period analyzed, suggesting that these processes likely do not contribute to
any overall changes in synapse density that occur during development. However, splitting
and merging of presynaptic components likely plays at least some role in refining circuits
over time. The extent to which splitting and merging of existing synaptophysin-tdtomato
structures contributes to this refining process, as well as the mechanisms responsible for
these phenomena remain to be determined.
It should be noted that spatial resolution is limited in the axial (Z) dimension due
to the point spread function of light detected through the objective. This could lead to
false detection of merging events, if one synaptophysin-tdtomato puncta moves directly
above or below another puncta and remains in close proximity to that puncta. It might
also lead to lower detection of splitting events, if a single puncta splits into two distinct
puncta that separate in the axial direction and do not move away from each other.
Resolution of the lateral (X, Y) dimensions is better than that of the axial dimension.
However, small distinct puncta that are in close lateral proximity to each other might be
detected as a single larger puncta or larger puncta that split but do not move away from
each other would not be detected. Again, this would lead to an overrepresentation of
merging events and an underrepresentation of splitting events in our data.
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Figure 4.3: Synaptophysin-tdtomato puncta both split and merge in vivo. (A) An
example of two synaptophysin-tdtomato puncta (red) merging over the course of minutes
within the intact mouse brain. (B) The percent of synaptophysin-tdtomato puncta that
merge and split is similar over the same developmental time-course, suggesting that these
processes do not account for major changes in synapse number at this developmental
stage. However, circuit refinement is a likely consequence of these dynamics.
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In addition to splitting and merging, a small population of synaptophysin-
tdtomato puncta was quite mobile within the cortex. These puncta were manually tracked
via Imaris software and subjected to analysis via custom-written MATLAB programming
(Figure 4.4A). Both instantaneous velocity and puncta displacement were measured for
mobile synaptophysin-tdtomato at each developmental time point. At the earliest
timepoint (P13-P15), the average instantaneous velocity was 0.141m/s, +/- 6.2x10-3
mm/s (Figure 4.4B, n = 35 movements), while it was slightly but significantly lower
during the two subsequent time points, indicating a reduction of rapid movement for the
most mobile puncta (Figure 4.4B, for P16-P19, inst. vel. = 0.129m/s, +/- 3.3x10-3
, n =
97 movements, p = 0.040 (compared to P13-15); for P20+, inst. vel. = 0.124m/s, +/-
7.9x10-3
n = 13 movements, p = 0.044 (compared to P13-15)). The average total distance
moved per puncta (normalized for length of the individual time-lapse image) was also
decreased with developmental age as this value was significantly lower at the latest time
point (Figure 4.4C, at P20+ distance moved = 0.15 m/puncta/min, +/-
0.045m/puncta/min, n = 8 puncta), compared to the two earlier time periods (Figure
4.4C, P13-15 distance moved = 0.70 m/puncta/min, +/- 0.13m/puncta/min, n = 16
puncta, p = 0.0016 (compared to P20+); P16-19 distance moved = 0.68m/puncta/min,
+/- 0.063m/puncta/min, n = 63 puncta, p = 4.8x10-5
(compared to P20+)). These data
taken together suggest that presynaptic terminals in the developing cortex become less
dynamic as time progresses.
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Figure 4.4: Instantaneous velocity and mean displacement decrease as development
progresses. (A) Series of time-lapse images showing movement of two different
synaptophysin-tdtomato puncta over time (white arrow, red arrowhead). (B)
Instantaneous velocity of moving puncta decreases slightly as development progresses.
(C) Normalized distance moved per minute also decreased at later developmental time
points. Scale bar = 10m, * = p-val < 0.05, ** = p-val < 0.005
111
Materials and Methods
Contributions
Time-lapse two-photon imaging was performed by TAE while tracking analysis with
Imaris software and MATLAB programming was performed by LADB.
Virus injection
LSL-synaptophysin-tdtomato homozygous mice were obtained from Jackson
Laboratories (stock #: 012570 (tdtom), Bar Harbor, ME, USA) and bred to obtain pups
with the flox-stop synaptophysin-tdtomato gene. For some experiments, these mice were
cross-bred with Thy-1-YFP mice (Jackson Laboratories, Bar Harbor, ME, USA). At
postnatal day 2-4, pups were anesthetized via hypothermia, after which AAV1-hSyn1-
Cre-IRES-GFP virus was injected into the right hemisphere of the cortex via Hamilton
syringe. Virus was created by the Vector Core at the University of Pennsylvania
(Philadelphia, PA, USA). Pups were then warmed by hand before being placed back into
their original cage.
Cranial window
At P10, pups that previously had undergone viral injection were anesthetized via
inhalation of approximately 1-3% isoflurane solution vaporized in pure O2 dispensed at a
rate of 1.5L/min. Pups were then given a subcutaneous injection of dexamethasone
(0.1mg/kg) and carprofen (5mg/kg) to prevent post-surgery edema and pain, respectively.
The head was then stabilized via stereotaxic instrument (Stoelting Co., Wood Dale, IL,
USA) before removing the skin on top of the head to expose the skull. A surgical
microdrill (Fine Science Tools, Foster City, CA, USA) was then utilized to gently cut
112
around and remove a section of the skull over the left hemisphere to expose the brain.
During drilling and after removal of skull piece, the surgical region was constantly
submerged in sterile saline in order to avoid inflammation during the procedure and to
keep the skull cool during drilling. Any minor bleeding that occurred was remediated by
gentle application of GelFoam (Baxter, Deerfield, IL, USA) to the affected area. Once
any bleeding was stopped, GelFoam was removed and a 5mm glass coverslip was placed
over the exposed region of the brain. The coverslip was attached to the remainder of the
skull via Vetbond glue (3M, Saint Paul, MN, USA) and Ortho-Jet self-curing dental
acrylic (Lang Dental Manufacturing, Wheeling, IL, USA). Acrylic was mounded around
the outside of the coverslip to create a well for the water immersion objective that was
used for imaging through the cranial window. In addition, a stabilization bar was
mounted in the acrylic. After surgery, mice were removed from anesthesia to recover on a
warm pad before being returned to their home cage.
Two-photon confocal imaging
To visualize synaptophysin-tdtomato, mice with cranial windows were anesthetized via
isoflurane (dose) and attached to a stereotaxic instrument via the stabilization bar that
was mounted in the dental acrylic upon cranial window surgery. These mice were then
transferred to a custom built heated chamber kept at 37oC that surrounded the objective of
the microscope. Isoflurane anesthesia was maintained via dedicated lines installed in the
environmental chamber. A Leica SP5 confocal microscope fitted with a 16W Ti/Sapphire
IR laser (Chameleon, Coherent, Inc., Santa Clara, CA, USA) and a 20x Leica HCX-APO-
L, N.A. 1.0 water immersion objective was used for excitation and imaging at
wavelengths tuned between 840 and 977nm. The tip of the objective was submerged in
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water that was pooled in the dental acrylic mound over the cranial window. The water
column was maintained by dedicated water lines included in the environmental chamber.
Time-lapse imaging was performed at a rate of 1 image per 30 seconds with 30 seconds
of rest in between images. Individual images consisted of z-projections with a step-size of
1.5m. After imaging, mice were allowed to recover in a warmed area before being
returned to their home cage.
Perfusion, sectioning, immunohistochemistry
Mice were deeply anesthetized via isoflurane inhalation and then transcardially perfused
with 1x PBS followed by 4% paraformaldahyde. Brains were then removed and drop-
fixed in 4% paraformaldahyde for 1 hour before being transferred to 30% sucrose
solution for cryoprotection. After brains were sunk in the sucrose solution, they were
frozen in dry-ice cooled 2-methylbutane and stored at -80oC until sectioning. Prior to
sectioning, tissue was mounted in OCT Compound (Tissue-Tek, The Netherlands) and
frozen on dry ice. Brains were then sectioned via Leica CM1850 cryostat at thicknesses
of either 10-15um (for antibody labeling) or 40um (for sections without antibody
labeling). Immunohistochemistry was then performed on thin (10-15um) sections
utilizing rabbit anti-synapsin primary antibodies (Synaptic Systems, Goettingen,
Germany) with anti-rabbit secondary antibodies conjugated to Alexa Fluor 488. Sections
were then mounted with fluoromount containing DABCO (Sigma, St. Louis, MO, USA)
and imaged via a C1 Plus confocal system on a Nikon Eclipse Ti-E microscope utilizing
a 20x Nikon Plan Apo 0.75NA objective.
Image analysis and statistics
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All synaptophysin-tdtomato puncta were manually tracked via Imaris software.
Movement was analyzed via custom written programs in MATLAB. Significance was
determined via Wilcoxon rank sum test unless otherwise indicated. All error bars are
S.E.M. unless otherwise noted.
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Chapter 5: Discussion
In Chapter 2 we hypothesized that trafficking of STVs and PTVs is coordinated
even prior to synapse assembly, which could facilitate rapid assembly. By performing
live multi-channel fluorescence confocal imaging in neurons expressing both
synaptophysin-mRFP and GFP-bassoon, we were able to record and analyze the
movements of STVs and PTVs simultaneously in the same neuron. The results indicate
that STVs and PTVs are coordinated during transport and before stabilization since PTVs
and STVs move together within the axon. We also find that STVs and PTVs pause at the
same sites within the axon, particularly when the other type of vesicle is also paused at
that site. This attraction of STVs and PTVs to the same pause sites is not mediated by a
direct interaction between STVs and PTVs since reducing the density of PTVs in the
axon yielded only small changes in STV pausing. In summary, the data support a model
of synapse formation in which STV and PTV trafficking is coordinated even prior to axo-
dendritic adhesion. This coordination includes coincident stopping of STVs and PTVs at
predefined sites of synapse formation, independent of a direct interaction between STVs
and PTVs.
Is STV and PTV transport coordinated?
Although it is clear that both PTVs and STVs need to be recruited to the same
sites in order for pre-synaptic terminals to develop, it is not immediately apparent as to
how they both arrive at the same destinations. Previously, STV and PTV dynamics have
been imaged only separately, leaving it unknown whether they are transported together
(47, 48, 50, 51, 53, 58-60, 210, 250). Recent work showed that clear, synaptic vesicle
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protein-containing vesicles and dense-core, PTV-like vesicles can be seen apparently
tethered together in electron micrographs of young neurons (49). This was the first
indication that PTVs and STVs might be transported together, but it remained unclear
whether these vesicle aggregates correspond to either STVs and PTVs being transported
together or paused together or a later stage in synapse development. Our analysis
indicates that a sizable portion of STVs and PTVs move with each other and spend the
majority of their time together. STVs and PTVs often moved separately before or after
moving together, suggesting that STVs and PTVs can move with each other while also
maintaining their separate identities. This observation indicates that, although it is
possible that some STVs contained molecules of GFP-bassoon and/or a portion of PTVs
contained molecules of synaptophysin-mRFP, missorting of STV and PTV marker
proteins cannot account for the observed co-transport of STVs and PTVs. Coordinating
STV and PTV movement could represent a mechanism for rapid synapse development,
since a full complement of synaptic proteins could immediately be delivered to a
potential synaptic site. It will be important in the future to determine whether co-
transport is mediated by a direct interaction between STVs and PTVs or by other
mechanisms.
Previously, it was shown that STV pause sites are preferred sites of synapse
formation (51), raising the question of whether PTVs also tend to pause at these same
sites of synapse formation. By labeling both STVs and PTVs within the same axon, we
were also able to compare spatial and temporal properties of pausing for both types of
vesicles. STV and PTV pausing were qualitatively and quantitatively similar. Like
STVs, multiple PTVs paused at the same sites, and the same PTVs returned repeatedly to
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a given site. The frequency of pausing and duration of pauses for STVs and PTVs were
also similar. Importantly, STVs and PTVs paused at the same sites within the axon. This
implies that PTVs also pause at predefined sites of synapse formation. The propensity of
STVs and PTVs to preferentially pause at the same sites within the axon suggests that
there is a mechanism that recruits both types of vesicles to the same site at the same time.
These same mechanisms could be utilized by axons to recruit the necessary proteins
during synapse formation.
What causes STV and PTV pausing?
Our data indicate that PTVs preferentially pause at sites where an STV is present.
Likewise, STVs preferentially pause at sites where a PTV is present. There are at least
three potential hypotheses which could account for this phenomenon (Figure 5.1). First,
pausing of STVs and PTVs at the same sites could be a result of STVs and PTVs
travelling together and consequently being recruited together. Second, it is possible that a
paused vesicle can interact with and stabilize a moving vesicle. In this scenario, a local
signal could cause one type of vesicle to pause at a specific site within the axon. That
paused vesicle could then interact with other vesicles and cause them to pause at the same
site. Third, STVs and PTVs could be independently attracted to a common signal within
the axon. In this case, both STVs and PTVs would respond to that signal by preferentially
pausing at the site of the signal, regardless of the presence of other vesicles. This, in turn,
would increase the probability that either vesicle was present at the site of the signal and,
therefore, the chance STVs and PTVs are simultaneously paused at the same site. Each
hypothesis represents a viable mechanism through which both synaptic vesicle and active
zone proteins could be recruited to the same site in the axon.
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Our data are most consistent with the hypothesis that STVs and PTVs are
recruited to the same sites at the same time as a consequence of responding to a common
local signal. The first mechanism – simultaneous recruitment of STVs and PTVs that are
transported together – cannot by itself explain coincident pausing since in many cases the
STVs and PTVs that paused together paused sequentially rather than simultaneously (e.g.
see Figure 2.3A, 2.3B-1 and 2.3B-3). To distinguish between the remaining two
mechanisms, we reduced the density of PTVs within the axon, using a dominant negative
construct corresponding to the syntaxin-binding domain of syntabulin. If PTVs interact
with STVs to influence STV pausing and recruitment to a given site, then decreasing
PTV density should substantially alter STV pausing. However, in SBD-transfected axons,
STVs displayed only small differences in pause duration and no change in the frequency
of pausing when compared to STVs in control axons. Although differences in pause
duration and velocity were statistically significant, the relatively subtle change in STV
pausing suggests that PTVs exert only a small influence on STV pausing. This argues
against the second hypothesis, in which STV and PTV pausing is exclusively controlled
by an interaction between STVs and PTVs. However, based on the data, it is still possible
that STVs and PTVs respond to both local signals and one another.
It will be important in the future to identify the signals that recruit STVs and
PTVs. It is not yet known which signals cause vesicles to pause. Potential signals include
calcium, phosphorylation or small GTPase activity. Although it was previously shown
that calcium could increase the duration of pausing, the probability of pausing was
unchanged by reducing intracellular calcium (51), suggesting that calcium alone may not
be responsible. Phosphorylation is known to regulate the activity of microtubule motor
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proteins and their association with microtubules or vesicles (101, 251-254), raising the
possibility that phosphorylation of motor proteins is the ultimate mechanism by which
local signals cause STVs and PTVs to stop their transport. Small GTPases are well-
known for their ability to control vesicle targeting (255-257), and recent work has shown
that in C. elegans, arl-8, an Arf-like small G-protein, controls where along the axon
synaptic proteins aggregate (95).
Why don’t all vesicles pause at any given site?
Although the majority of STVs and PTVs that encounter pause sites will pause at
them, many vesicles ignore the pause sites and continue their movements. This is even
true for stopping and recruitment that occurs in response to synaptogenic adhesion or
axo-dendritic contact (51). It is not clear why. One possibility is that there could be a
limited number of “docking sites” at each pause site. This seems unlikely since some
vesicles pass by even while others can still pause. Alternatively, STVs and PTVs could
exist in multiple states, at least one of which is unresponsive to local signals that cause
pausing. PTVs and STVs display heterogeneity in apparent size, velocity, and pausing
properties, which has been noted here and elsewhere (47, 48, 51, 53, 58, 60, 73). These
apparent differences may correlate with structural differences (49). In general,
smaller/dimmer vesicles appear more likely to move quickly and pass by pause sites
without pausing. Perhaps these apparently smaller vesicles are unable to respond to the
cues that induce pausing. Such a state could be a result of differences in their motors or in
other proteins associated with the STVs or PTVs. The idea of STVs and PTVs as
heterogeneous sub-populations will be an interesting topic for further exploration.
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A model of coordinated transport
For synapses to form, all of the components of presynaptic terminals must be
delivered rapidly to the site of synapse formation. Despite its importance, we are only
beginning to understand how this occurs. Here we have proposed a model for presynaptic
terminal assembly in which trafficking of synaptic vesicle and active zone proteins is
coordinated even prior to axo-dendritic contact. This coordination occurs through a
combination of co-transport, perhaps in aggregates of vesicles tethered together (49), and
co-pausing in response to local signals. This coordination then can facilitate presynaptic
terminal assembly and contribute to the rapid recruitment of synaptic components that
has been consistently observed. The same signals that induce co-pausing prior to
transport may act down-stream of synaptogenic adhesion to simultaneously attract STVs
and PTVs to sites of synapse assembly.
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Figure 5.1: Models for coordination of recruitment of STVs and PTVs to the same
place at the same time. STVs and PTVs could be simultaneously attracted to a given site
by (1) attraction of STVs and PTVs that are co-transported; (2) sequential recruitment via
a physical interaction between STVs and PTVs; and (3) simultaneous but independent
recruitment in response to a common signal. Our data are most consistent with the third
model.
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What are the dynamics of presynaptic protein recruitment at developing terminals?
In Chapter 3, live, time-lapse confocal imaging of fluorescently-tagged proteins
was performed at developing presynaptic terminals which were artificially induced
through the interaction of trans-synaptic adhesion molecules. By imaging presynaptic
proteins at newly induced synapses over the course of minutes and hours, we were able to
determine that presynaptic recruitment is much more labile than previously thought. In
addition, live imaging of developing synapses in axons co-expressing synaptophysin-RFP
and neurexin-GFP established the timecourse and dynamics through which neurexin
mediates it synaptogenic signal. Furthermore, tracking STVs and PTVs outside of
synaptogenic regions strongly suggested that these components are trapped at developing
synapses, rather than actively recruited to them.
Presynaptic proteins are trapped at, but not attracted to, developing synapses
Presynaptic terminal formation is initiated by contact between axons and
dendrites, which leads to interactions between trans-synaptic adhesion molecules,
recruitment of presynaptic proteins and subsequent organization of presynaptic terminals
(Figure 5.2). It remains unclear how trans-synaptic signaling recruits presynaptic
proteins. On one hand, trans-synaptic adhesion could actively attract transport vesicles to
sites of axo-dendritic contact. Alternatively, trans-synaptic adhesion could establish sites
in the axon where synaptic proteins become trapped and organized into a presynaptic
terminal, without actively attracting transport vesicles.
Our results favor the “trapping” model (Figure 5.2, step 2, top). STVs and PTVs
did not preferentially move towards sites of trans-synaptic signaling, and their transport
123
was minimally altered by trans-synaptic adhesion. These findings suggest that trans-
synaptic adhesion does not actively attract transport vesicles into developing presynaptic
terminals. Instead, these data support the hypothesis that trans-synaptic adhesion
establishes sites in the axon that capture and stabilize available presynaptic protein.
Importantly, STVs were not attracted to SynCAM-1 or both neuroligin-1 and SynCAM-
1, and STV movement was unaffected by this combination of trans-synaptic signaling.
Therefore, assembly of synaptic proteins at developing presynaptic terminals via
establishment of sites of stabilization (rather than active attraction) appears to represent a
general mechanism that is conserved across synaptogenic pathways.
SV proteins preferentially accumulate at predefined sites that are intrinsic to the
axon (51). Although it is not known what stabilizes synaptic proteins at these specific
sites, STVs frequently pause at these sites prior to synaptogenesis. Trans-synaptic
adhesion might facilitate recruitment by stabilizing paused STVs (5, 51, 136). It is
important to note that STVs carry a variety of synaptic vesicle and some active zone
proteins (53), so it remains unclear which proteins mediate any interactions (direct or
indirect) between STVs and neurexin or SynCAM. ARL-8, which is transported with
STVs in C. elegans, has recently been shown to control aggregation and delivery of SV
and AZ proteins to appropriate sites of synaptogenesis (95, 96). Synaptogenic adhesion
could act through regulation of ARL-8 or a similar pathway to stabilize paused STVs.
The trapping model predicts that the supply of synaptic proteins in the axon
would determine the probability that synaptic proteins encounter these sites and,
therefore, levels of trapping and recruitment. Consistent with this, synapse formation is
enhanced by increasing the number of mobile STVs in the axon (160, 258), and synaptic
124
protein recruitment to poly-D-lysine coated beads preferentially occurs in axons with
high levels of protein expression and mobility (116). Interestingly, disruptions in
transport have been linked to defects in synapse formation and function (95, 96, 102, 111,
259, 260) and associated with intellectual disability in humans (261).
Synaptic proteins are labile at developing synapses
While there have been studies of the stability of new synapses (215, 262), it
remained unclear how stable synaptic proteins are at synapses after their initial
recruitment. Here, we showed that enrichment of SV proteins at individual trans-synaptic
signaling sites is highly labile: although SV proteins remained enriched at these sites, the
degree of enrichment rapidly increased and decreased, oftentimes varying over the course
of minutes. Similar fluctuations in synaptic protein levels have been observed at mature
synapses, where changes in levels of synaptic vesicles and active zone proteins have been
directly linked to changes in synapse function (263-265). At mature synapses, lability in
presynaptic protein levels arises through sharing of presynaptic components between
adjacent synapses (218-221, 223-225, 266, 267), and individual shared SVs can be
functionally integrated into the synapse within minutes after their arrival (223, 267). Our
results suggest that this dynamic arrival and departure (fluctuation) of SVs at synapses
occurs at the earliest stages in synapse development. It remains to be seen whether SVs at
developing synapses can rapidly become incorporated into the synapse and fuse with the
plasma membrane in response to action potentials. However, STVs can fuse with the
plasma membrane upon neuronal depolarization (47, 51, 58) and most likely release
glutamate at these sites when they do (51). Since glutamatergic signaling regulates the
accumulation of presynaptic proteins at developing synapses (268-270), varying the level
125
of presynaptic proteins at new excitatory synapses could alter glutamate release and
contribute to feedback or auto-regulation of synapse maturation.
We observed both gradual and sudden, dramatic changes in the amount of
synaptic proteins recruited. These two types of fluctuations in recruitment may represent
distinct features of presynaptic growth and stabilization. The sudden appearance of
synaptophysin at the site of trans-synaptic adhesion indicates that presynaptic proteins
can be recruited and stabilized at contacts on the order of seconds. This is striking
considering recent evidence that dendritic spines can form in seconds in response to
glutamate uncaging (271). Together, these data suggest that complete synapses can form
extremely rapidly. In addition, the immediate loss of synaptophysin puncta that we
observed suggests that developing synapses can be eliminated equally quickly. Finally,
the gradual accumulation or loss of synaptophysin that occurred at many contacts may
correspond to changes in presynaptic maturation or strength (272).
A model of neuroligin-neurexin mediated presynaptic protein recruitment
It has been proposed that synaptogenesis occurs in a 2-step model: postsynaptic
clusters of neuroligin induce neurexin clustering, then clustered neurexin seeds the
recruitment of presynaptic proteins (124). Our data support the hypothesis that neurexin
clustering is critical for stable synaptic protein recruitment. However, we also found that
neurexin clustering and synaptophysin recruitment appeared simultaneously. Although
the two steps of the model could occur sequentially but very rapidly, an intriguing
alternative explanation is that a cue within the axon cooperatively contributes to both
neurexin clustering and STV accumulation. Consistent with this idea, recent work in
Drosophila showed that neurexin clustering is mediated by axonal syd-1, and that this
126
process is critical for the synaptogenic ability of neurexin (125). Syd-1 acts upstream of
syd-2/liprin-α, a protein linked to transport of STVs and initiation of active zone
formation (77, 196-198, 203, 273, 274). It will be important in the future to determine
whether similar molecular mechanisms link neurexin clustering to STV stabilization in
mammals. Although mammalian orthologs of syd-1 do not contain the requisite PDZ for
neurexin organization, mammalian syd-1 (mSYD1A) has recently been shown to both
regulate SV protein accumulation during synaptogenesis and interact with liprin-α2
(206). Interestingly, in Drosophila, feedback from the postsynaptic partner also
contributes to presynaptic assembly and stabilization (125). We found that the presence
of the postsynaptic scaffolding molecule S-SCAM in neuroligin-1-expressing HEK-293
cells increased the degree of neurexin clustering in the axon, consistent with analogous
cooperative pre-and postsynaptic mechanisms existing in mammals.
While many proteins that interact with the cytoplasmic domain of neurexin have
been thought to be attractive candidates for mediating neurexin’s synaptogenic signaling
within the axon, recent work has revealed that the intracellular tail of neurexin is
completely dispensable for its synaptogenic properties (217). This suggests that neurexin
utilizes unidentified cis interactions with axonal transmembrane proteins to signal
intracellularly. It will be important to identify the cis-interacting neurexin partner(s) and
its cytosolic interactions in order to understand neuroligin/neurexin-mediated synapse
formation.
Here, we have focused on synapse assembly down-stream of neuroligin-neurexin
signaling, the prototypical synaptogenic adhesion pair. Additional factors also play a role
in regulating protein levels at presynaptic terminals, including local actin polymerization
127
(117, 234, 235, 275), signaling through BDNF/TrkB (258, 276-279) and NMDA
receptors (270), altering VGLUT1 expression (268, 269), and through other trans-
synaptic adhesion molecules not addressed in our study (136). These signals could
regulate neurexin clustering, function down-stream of neurexin clustering, or operate
through parallel mechanisms. Either way, these mechanisms likely work in concert to
dynamically regulate synapse development and function.
A model for dynamic presynaptic protein recruitment to a developing synapse
Our results strongly support a model of synapse formation where presynaptic
proteins are trapped at developing presynaptic sites, rather than actively attracted to them
(Figure 5.2, steps 1-2). Trapping tends to occur at sites of neurexin clustering, but not at
sites of diffuse neurexin recruitment, supporting the hypothesis that the spatial
arrangement of neurexin is critical for neurexin/neuroligin induced synaptogenesis.
Neurexin clustering and SV protein recruitment occur simultaneously (Figure 5.2, step 3).
Finally, at individual nascent presynaptic terminals, SV protein levels can vary within
minutes to hours, and these levels correlate with the amount of clustered neurexin. These
fluctuations might, in turn, affect later stages of synapse formation, maturation and
function.
128
Figure 5.2: Model of synaptic vesicle protein transport vesicle (STV) recruitment to
a developing presynaptic terminal. (1) Axo-dendritic contact induces diffuse neurexin
recruitment to the contact site. (2) STVs are not attracted to this site (bottom). Instead,
they are trapped as they encounter this site during normal axonal transport (top). (3) As
STVs become trapped, neurexin simultaneously clusters, resulting in the formation of a
stable site of synaptic protein accumulation.
129
In vivo synaptic dynamics
A number of non-mammalian experimental models have been developed to
address presynaptic protein recruitment and terminal formation within intact organisms
(96, 126, 280-289). In mice, two-photon confocal imaging has been used to analyze
postsynaptic spine formation and elimination within the intact brain by tracking
morphological changes of dendritic spines in GFP-filled cells (290-292). Presynaptic
bouton dynamics have also been evaluated in the mouse brain by identifying and
studying axon swellings in GFP/RFP/tdtomato-filled cells (293-296). However, the
identification of real presynaptic boutons is somewhat arbitrary as they are difficult to
distinguish from non-synaptic axonal swellings (297). Furthermore, smaller, more
dynamic synapses might be undetectable as axonal swellings.
In Chapter 4, we address these concerns though two-photon imaging of
synaptophysin-tdtomato in the brain of living mice through cranial windows. We find
that these fluorescently-tagged presynaptic proteins properly localize to synapses in the
brain, making it an excellent tool to study how presynaptic structures change over time.
Live, time-lapse imaging indicated that components such as synaptophysin-tdtomato in
many presynaptic terminals within the intact brain change over minutes via splitting,
merging, or wholesale movement. In addition, these experiments have shown that the
frequency and extent of these dynamic processes are altered over the course of
development, suggesting a possible functional role in the formation of circuits within the
intact brain.
130
A number of questions remain in regards to these dynamic presynaptic processes
which can be addressed by the type of in vivo imaging described here. For example, what
are the functional consequences of splitting, merging, or moving a presynaptic terminal
for the circuit? These experiments also only addressed the dynamics of presynaptic
terminals over approximately one week of development. Do these processes occur at
earlier timepoints? Do they occur and/or are they important in the aging brain? In
addition, the live imaging of synaptophysin-tdtomato was performed mainly in layer I of
the cortex. Are the dynamics of presynaptic terminals different in different layers?
Furthermore, are there any differences in presynaptic dynamics in different cell types
and/or cortical areas? While mice in these experiments were anesthetized, it is most likely
possible to perform similar experiments in awake behaving mice (298). It would,
therefore, be interesting to determine whether environmental stimuli affect presynaptic
dynamics. Finally, it will be critical to determine whether these dynamics are altered in
disease states.
Future outlooks – directions for future investigations
N-cadherin: trans-synaptic facilitator
An informative example of how elucidating the interactions between different
trans-synaptic adhesion systems can lead to a more complete understanding of
downstream synaptogenic signaling is illustrated by n-cadherin. N-cadherin, a
homophilic trans-synaptic adhesion protein, was one of the first trans-synaptic proteins
identified and was immediately hypothesized to be important for synapse formation. It
has since been shown that n-cadherin and its intracellular signaling through -catenin, -
131
pix, and scribble are important for localizing SVs to the presynaptic terminal through F-
actin polymerization (160, 161, 235). N-cadherin is also recruited to newly formed
synapses in zebrafish on the same time course as STVs (283). However, the importance
of n-cadherin for synaptogenesis decreases as neurons mature (161). In addition, synapse
number actually increases in -catenin knockout mice (160), possibly due to enhanced
SV distribution within the axon (258). Furthermore, n-cadherin by itself cannot induce
presynaptic terminal formation, as axonal contact with a non-neuronal cell expressing n-
cadherin does not lead to presynaptic terminal formation at the contact site as it does with
other post-synaptic adhesion proteins such as neuroligin and SynCAM (128, 157).
Despite this, n-cadherin interacts with other trans-synaptic adhesion proteins to
promote their function. For example, n-cadherin mediates the synaptogenic capability of
neuroligin in dendrites, most likely by enhancing neuroligin clustering postsynaptically
through intracellular interactions with -catenin and the synaptic scaffolding molecule
(S-SCAM) (216, 233, 299). In addition, the n-cadherin/-catenin complex interacts with
members of the leukocyte common antigen-related family receptor protein tyrosine
phosphatase (LAR-RPTP) family of proteins, which are important for the development
and organization of presynaptic terminals (300, 301). Also, n-cadherin is a substrate for
PTP, which can form a trans-synaptic adhesion complex with postsynaptically localized
TrkC to mediate synapse formation (169, 302). More work is necessary to characterize n-
cadherin’s interactions with other axonal trans-synaptic adhesion molecules. However, by
studying multiple trans-synaptic adhesion complexes in concert, we are beginning to
understand the complex and intertwined set of instructions that underlie synapse
formation.
132
Actin
Regulation of the axonal actin cytoskeleton may be an important step in
presynaptic terminal assembly down-stream of synaptogenic adhesion. Several synaptic
adhesion molecules control assembly of a localized network of F-actin at nascent
synapses, and formation of this cytoskeletal structure is required for synapse formation
(235, 236, 275, 303, 304). Actin depolymerization during early stages of development
leads to reduced synapse size and number (234), and various F-actin regulators play a
role in synapse development in mammals, Drosophila, and C. elegans (236, 305-309).
Importantly, the precise role of the actin cytoskeleton in presynaptic development is
beginning to be unraveled. It has long been known that F-actin within the presynaptic
terminal organizes synaptic vesicles and may localize them near the active zone
membrane. Prior to synapse assembly, depolymerization of actin alters the mobility of
STVs and their pausing behavior at sites of eventual synapse formation (51). F-actin can
also interact with active zone proteins and control active zone protein assembly at
synapses (275). In turn, active zone proteins, such as piccolo, can promote the formation
of presynaptic F-actin structures (193).
New technologies for studying synaptogenesis
It is clear that an important goal of developmental neuroscience is to understand
the mechanisms of synaptogenesis, but it is equally clear that many large gaps remain in
our understanding of the process of synapse formation. In many ways, the nature of
synapse formation has made this a difficult problem to attack. For example, organisms
need synapses to survive, limiting what can be discovered using traditional genetic
screens. In addition, observing the events of synapse formation can be challenging
133
because of (1) the unpredictable nature of when and where en passant synapses form in
the brain and (2) the rapid assembly of individual presynaptic terminals once the process
is initiated at any given axo-dendritic contact. Moreover, synapse elimination occurs
concurrently with synapse formation, making it impossible to determine whether
manipulations have specifically altered synapse assembly (as opposed to elimination) in
fixed tissue. Therefore, approaches such as fast, multi-channel time-lapse imaging are
necessary, in order to examine the time course of changes at individual synapses.
Application of recent advances in time-lapse imaging to the study of synapse
assembly should propel the field forward. For example, time-lapse imaging has
traditionally relied on the use of exogenously expressed proteins bearing fluorescent tags,
such as GFP, but exciting new methods have been developed that allow fluorescence
imaging of endogenous proteins (237, 310). In addition, the vast majority of studies
applying time-lapse imaging to study mammalian synaptogenesis have been performed in
culture. While it is likely that the core mechanisms of synaptogenesis are conserved in
vitro, it will be important to verify this with live in vivo imaging of presynaptic terminal
formation. Pioneering studies have examined presynaptic development in non-
mammalian models and at the mammalian neuromuscular junction (96, 126, 280-288);
however, presynaptic terminal development and synaptic protein recruitment will need to
continue to be investigated within the mammalian brain utilizing experiments like those
described in Chapter 4.
Another technical advance with promise to dramatically improve our
understanding of the mechanisms of synapse assembly is the development of high-
throughput genetic screens in mammalian neurons. For decades, elegant genetic screens
134
in Drosophila and C. elegans have provided insight into the molecular mechanisms of
synapse formation. Recently, screens have been used to identify molecules involved in
mammalian synaptogenesis, as well (169, 214, 311-313). These screens have been
particularly useful for identifying cell surface receptors that play a role in synaptogenesis.
Future screens should be helpful in identifying intracellular signals that orchestrate
synapse assembly. In addition, progress in cultivating neurons and organoids from human
induced pluripotent stem cells suggests such screens could be performed on human
neurons in the near future (314-316).
Conclusion
Synaptogenesis is a continuous process that occurs trillions of times over the
course of a human life, and its disruption can lead to devastating neurological and
psychiatric consequences. Although significant advancements into understanding
presynaptic development have been made, much more work is required. Critical to this
endeavor will be investigations into how all of these signaling mechanisms interact with
one another and lead to the dynamics observed in presynaptic development. Additionally,
insights into how disease-associated mechanisms affect the coordination of these
processes are needed. Ultimately, this will lead to a better understanding of how circuits
are established in the brain and how they are altered in disease.
135
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