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Uncovering cell type-specific roles of proteins involved in glutamatergic synapse regulation

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Uncovering cell type-specific roles of proteins involved in glutamatergic synapse regulation

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Content Uncovering cell type-specific roles of proteins involved in glutamatergic synapse regulation
by
Anna Pushkin
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(NEUROSCIENCE)
May 2024
Copyright 2024 Anna Pushkin

ii
Dedication
For Natalia, Alexander, Lika, and Nico

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ACKNOWLEDGEMENTS
The work in this dissertation would not have been possible without my graduate advisor,
Dr. Bruce Herring. Bruce, your passion for science is infectious. Thank you for always listening
to my ideas and helping me to grow as a scientist. You are a true mentor. To all members of the
Herring lab, past and present – you have made this journey an immense joy. I am eternally
grateful to you all for making work a fun and hilarious place to be. Special thanks to Chen and
Yuni, my snack-enabling work moms. Thank you for always believing in me, especially when I
did not believe in myself. I will cherish the memories of cookies and caffeine shared over 3PM
coffee hours.
I would like to extend my deepest gratitude to the members of my dissertation committee,
Dr. Dion Dickman and Dr. Karen Chang, for their helpful feedback and tremendous support. To
the earlier members of my guidance committee, Dr. Dave McKemy and Dr. Judith Hirsh, I
sincerely thank you as well. Your written prompts for my competency exam greatly improved
my technical writing and confidence in myself as a scientist. I am deeply grateful to my friends
and colleagues within NGP and the Neurobiology section as well: specifically, Jessica, Morgan,
Dawn, and Deanna.
Huge shout out to the 2018 NGP cohort, who made the transition to grad school a
memorable and joyous one. I still think fondly of our potlucks, study sessions, and shared meals.
Special shout out to Tanisha and Erin. This PhD truly would not be possible without you both.
Lastly, I would like to dedicate this to my biggest support system – my family. To my
parents, Alexander and Natalia. The sacrifices you made immigrating to a new country and
starting from scratch gave me the opportunities I have today. Your eternal love, support, and
willingness to listen have made me the human I am today. To Lika, the greatest older sister a

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person could ever ask for. Your fierce and caring nature, devotion, loyalty, and calm disposition
make you a rock for our family. To Sasha, the pup who teaches us all what unconditional love is.
And to Nico, the future of our little family. I thank you all.

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TABLE OF CONTENTS
DEDICATION .............................................................................................................................. ii
ACKNOWLEDGEMENTS.......................................................................................................... iii
LIST OF FIGURES ……............................................................................................................. vii
ABBREVIATIONS .................................................................................................................... viii
ABSTRACT ................................................................................................................................ ix
CHAPTER 1: INTRODUCTION ................................................................................................ 1
1.1 Glutamatergic Synapses ................................................................................. 1
1.2 The Hippocampus........................................................................................... 3
1.3 CA1 vs DG ..................................................................................................... 5
1.3 Different Molecular Regulators within DG.................................................... 8
CHAPTER 2: PROTEIN 4.1N PLAYS A CELL TYPE-SPECIFIC ROLE IN HIPPOCAMPAL
GLUTAMATERGIC SYNAPSE REGULATION ..................................................................... 12
2.1 Abstract .......................................................................................................... 12
2.2 Introduction .................................................................................................... 12
2.3 Methods .......................................................................................................... 14
2.4 Results ............................................................................................................ 21
Protein 4.1N is required for the structure and function of glutamatergic
synapses in DG granule neurons. .....................................................................21
4.1N knockdown does not perturb basal glutamatergic synapse function
in CA1 pyramidal neurons................................................................................25
Protein 4.1N’s C-Terminal Domain is not critical to its function at DG
granule synapses. ..............................................................................................26
Protein 4.1N’s FERM domain is required for synaptic AMPA receptor
function in DG granule neurons. ......................................................................28
2.5 Discussion ...................................................................................................... 31

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CHAPTER 3: THE ROLE OF THE 4.1 FAMILY OF PROTEINS IN HIPPOCAMPAL
GLUTAMATERGIC SYNAPSE REGULATION...................................................................... 46
3.1 Abstract .......................................................................................................... 46
3.2 Introduction .................................................................................................... 46
3.3 Methods .......................................................................................................... 49
3.4 Results ............................................................................................................ 53
4.1R knockdown does not perturb basal glutamatergic synapse function in
CA1 pyramidal neurons or DG granule neurons ............................................ 53
4.1B knockdown does not perturb basal glutamatergic synapse function in
CA1 pyramidal neurons or DG granule neurons ............................................ 54
Loss of 4.1N’s SAB domain augments synaptic AMPAR currents in DG
granule neurons ............................................................................................... 54
3.5 Discussion ...................................................................................................... 55
CHAPTER 4: CONCLUSIONS................................................................................................... 66
REFERENCES ............................................................................................................................ 70
APPENDIX: INVESTIGATING A POSTSYNAPTIC ROLE FOR NEUREXINS....................86
5.1 Introduction .................................................................................................... 86
5.2 Methods .......................................................................................................... 88
5.3 Results ............................................................................................................ 91
Knockdown of NRXN1α and NRXN1β (NRXN1 KD) does not perturb
basal glutamatergic synapse function in CA1 pyramidal or DG granule
neurons ............................................................................................................ 91
Knockdown of all neurexins (NRXN1, NRXN2, and NRXN3) does not
perturb basal glutamatergic synapse function in CA1 pyramidal or DG
granule neurons................................................................................................ 92
5.4 Discussion ...................................................................................................... 93

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LIST OF FIGURES
Figure 2.1: Protein 4.1N is required for glutamatergic synapse structure and function in DG
granule neurons ............................................................................................................................ 36
Figure 2.2: Protein 4.1N is not required for glutamatergic neurotransmission in CA1
pyramidal neurons ........................................................................................................................ 39
Figure 2.3: Protein 4.1N’s C-terminal domain is not required for glutamatergic synapse
function in DG granule neurons .................................................................................................. 41
Figure 2.4: Protein 4.1N’s FERM domain is required for maintaining synaptic AMPA
receptor function in DG granule neurons ................................................................................... 43
Figure 3.1 Conserved domain structure of the protein 4.1 family ............................................... 47
Figure 3.2 Hippocampal mRNA expression varies between members of the 4.1 family of
proteins (Figure adapted from Parra et al., 2000) ........................................................................ 48
Figure 3.3: Knockdown of protein 4.1R does not disrupt basal glutamatergic
neurotransmission in either CA1 pyramidal or DG granule synapses.......................................... 60
Figure 3.4: Knockdown of protein 4.1B does not disrupt basal glutamatergic
neurotransmission in either CA1 pyramidal or DG granule synapses.......................................... 62
Figure 3.5 Loss of protein 4.1N’s SAB domain augments synaptic AMPAR currents in DG
granule neurons ........................................................................................................................... 64
Figure 5.1: Domain structure of α- and β-neurexins ................................................................... 86
Figure 5.2: Knockdown of neurexin-1 does not disrupt synaptic transmission at CA1
pyramidal synapses or DG granule synapses............................................................................... 95
Figure 5.3: Knockdown of all neurexins does not disrupt synaptic transmission at CA1
pyramidal synapses or DG granule synapses............................................................................... 97

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ABBREVIATIONS
aCSF, artificial cerebrospinal fluid
AMPAR, α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor
CA1-3, cornu Ammonis 1-3
CNS, central nervous system
DG neurons, dentate granule neurons
DG, dentate gyrus
DIV, days in vitro
EC, entorhinal cortex
eEPSC, evoked excitatory postsynaptic currents
FERM, four-point-one, ezrin, radixin, and moesin
KD, knockdown
MAGUK, membrane-associated guanylate kinase
NMDAR, N-methyl D-aspartate receptor
PDZ, PSD-95/Discs large/Zona occludens-1
PNS, peripheral nervous system
PSD, postsynaptic density
P_, postnatal day _
RNAi, RNA interference
SAB, spectrin-actin binding
SEM, standard error of the mean
shRNA, short hairpin RNA

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ABSTRACT
The molecular mechanisms governing glutamatergic synapse structure and function are
vast and complex. Proper synaptic regulation is critical for healthy brain functioning. Historically,
most studies examining synaptic function have focused on Schaffer collateral-CA1 synapses and
extrapolated findings to be true for all synapses. Recent technological advances have begun to
uncover the immense functional, morphological, and molecular differences between synapse
subtypes. Despite this, more work needs to be done to truly understand and appreciate the extent
of these differences. 4.1N is a protein whose synaptic function has long eluded researchers,
despite its binding to multiple synaptic regulatory proteins. Chapter 2 details a novel, sitespecific, postsynaptic role for protein 4.1N in regulating perforant path – granule neuron synapses
of the dentate gyrus (DG). This role is partially dependent on 4.1N’s FERM domain. Our results
strongly suggest that 4.1N’s FERM domain plays a critical role in GluA1-containing AMPAR
trafficking to DG granule synapses. Chapter 3 introduces a second role for protein 4.1N in
synaptic regulation. Removal of 4.1N’s SAB domain dramatically increases synaptic AMPARmediated currents, highlighting the multifunctionality of 4.1N’s role in maintaining stable
AMPAR expression at DG granule synapses. Chapter 3 also investigates potential roles for 4.1R
and 4.1B, other 4.1 family members with localized hippocampal expression, in synaptic
regulation. Overall, my hope is that the work in this dissertation contributes to the growing body
of research uncovering the molecular specificity between synapse subtypes. Specifically, this
work further confirms perforant pathway synapses are governed by a unique set of molecular
regulators.

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CHAPTER 1: INTRODUCTION
1.1 Glutamatergic Synapses
Healthy brain functioning requires the ability to receive, store, and transmit mass amounts
of information. Neurons, specialized cells within the central nervous system, transmit information
between each other at synapses. Synapses are the sites of communication between neurons
(Lodish, 2000). Excitatory glutamatergic synapses ‘communicate’ through presynaptically
released ions binding to postsynaptically expressed ionotropic glutamate receptors. Among these
receptors, α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors (AMPARs) and Nmethyl D-aspartate receptors (NMDARs) play the largest known roles in excitatory
neurotransmission (Ozawa et al., 1998).
AMPARs are ionotropic transmembrane receptors composed of combinations of four
subunits (GluA1-GluA4), encoded by the genes GRIA1-GRIA4 (Hollmann and Heinenmann
1994; Traynelis et al., 2010). These ligand-gated channels are critical for basal synaptic
transmission, as their activation is fast and transient. They also undergo rapid recruitment during
synaptic plasticity (Collingridge et al., 2009). NMDARs are heteromeric ionotropic glutamate
receptors composed of GluN1, GluN2, and GluN3 subunits. (Cull-Candy et al., 2004). Synaptic
NMDARs are localized to the postsynaptic density (PSD). At mature synapses, NMDARs are
composed primarily of GluN1/GluN2A or GluN1/GluN2A/GluN2B assemblies. As with
AMPARs, the composition and number of NMDARs at synapses is a dynamic process based on
development, sensory experience, and activity (Lau et al., 2007).
NMDARs activate and deactivate at a far slower rate than AMPARs. This is partly
because they are both ligand- and voltage- gated ion channels. Glutamate binding from the
presynaptic cell to postsynaptic AMPARs triggers the influx of positively charged Na+ ions and

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subsequent depolarization into the postsynaptic neuron. Sufficient depolarization removes the
Mg2+ block on the NMDARs, allowing for the influx of Ca2+ into the cell. Ca2+ influx is essential
for synaptic plasticity and long-lasting changes in synaptic efficacy, such as LTD and LTP
(Collingridge et al., 1983). LTP, a rapid, long-lasting increase in synaptic strength, was
discovered in excitatory synapses at dentate granule (DG) neurons (Lomo, 1971; Bliss and Lomo,
1973). LTP occurring at Schaffer collateral-CA1 synapses represents the most widely studied
form of synaptic plasticity (Herring and Nicoll, 2016). To this day, LTP is believed to be one of
the main contributors to synaptic plasticity – and thus, learning and memory. LTD, a long-lasting
decrease in the efficacy of synaptic transmission, alters synaptic strength in the opposite direction
(Ito and Kano, 1982). Both AMPA and NMDA receptors have long been acknowledged for their
roles in these forms of synaptic plasticity (Dudek and Bear, 1992). When calcium flows into the
cell, it activates kinases such as CaMKII (Lisman et al., 2012). Activated CaMKII phosphorylates
the GluA1 subunit, which promotes synaptic integration of GluA1-containing AMPARs and
produces a rapid increase in the number of AMPARs at the synapse (Patterson et al., 2010). At
mature synapses, the number of AMPARs at a synapse correlates with spine size and synapse
strength (Matsuzaki et al., 2001).
NMDAR have long been viewed as molecular coincidence detectors of pre- and
postsynaptic firing (Bliss and Collingridge ,1993; Blanke et al., 2009). Activation of NMDARs
requires a sufficient depolarization as well as presynaptic glutamate release. This dual activation
requirement allows NMDARs to decode and integrate synaptic input, lending them a central role
in synaptic plasticity. NMDARs are thought to induce long term plasticity by modulating
AMPAR-mediated fast transmission (Malenka and Nicoll, 1993). Glutamate uncaging
experiments show that repetitive activation of NMDARs on a single spine is enough to produce a

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long-lasting increase in AMPARs from the same spine (Harvey and Svoboda, 2007; Matsuzaki et
al., 2004). In order to induce long lasting changes in synaptic transmission, both AMPARs and
NMDARs interact with a number of proteins that help traffic them into the synapse and regulate
their function. This dissertation will focus on the multifunctional, site-specific role of protein
4.1N in regulating glutamatergic synapses of the hippocampus.
1.2 The Hippocampus
Many pivotal discoveries into the molecular mechanisms governing glutamatergic synapse
structure and function have come from investigations into the hippocampus. Recognized for its
role in the spatial and temporal aspects of memory; the hippocampal formation has long been
considered to organize episodic memories in space (Eichenbaum et al.,1999).
To understand the function of the hippocampus, one must understand anatomical
projections within the hippocampus and beyond. The entorhinal cortex (EC) is the major source
of excitatory glutamatergic input into the hippocampus (Amaral and Witter, 1989). EC is a
polymodal sensory association area which receives projections from multiple cortical regions and
all sensory systems. Information flows from the EC via the axons of the perforant pathway (van
Strien et al., 2009). EC projects to all hippocampal subdivisions (DG, CA3, CA1….) and intrinsic
pathways involve successive projections from CA3 to CA2 and CA1 as well as from CA2 to CA1
(Gilbert and Brushfield, 2009). DG neurons send their axons (mossy fibers) to CA3. CA3 neurons
then send their axons to CA1 through the Schaffer collateral pathway. EC axons also terminate
directly in CA1, which contains pyramidal axons that are sent back to the deep layers of the EC.
While the basic hippocampal circuitry remains similar along the septotemporal axis, there
is substantial variation in the input projections between dorsal and ventral hippocampal regions.

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The dorsal hippocampus (DH) receives input from the dorsal EC which carries largely
visuospatial information (Dolorfo and Amaral, 1998). The ventral hippocampus (VH), on the
other hand, is innervated by the amygdala (Risold and Swanson, 1997). Classically, the DH is
associated with episodic memory and cognitive functions while the VH corresponds to stress and
emotion (Bliss and Collingridge, 1993; Fanselow and Dong, 2010). Lesions to the DH impair
performance on the Morris water maze, a spatial memory task, while VH lesions neither affect
nor exacerbate the deficit (Moser et al., 1995). Lesions to the VH, on the other hand, enhance
ulcers produced from cold and restraint stress (Henke, 1990).
Dorsal hippocampal neurons in rodents appear to encode highly specific location and
object-to-place combinations, whereas ventral hippocampal neurons appear to code for large areas
of space (Eichenbaum, 2017). It is becoming apparent, however, that dorsal vs ventral is not the
best way to differentiate function. While it is easier to anatomically divide the hippocampus this
way for lesion studies, doing so risks summation from disparate subfields and cell types to mask
individual effects. Historically, one way to disentangle the function of various subfields has been
to study neurons individually.
Studies into the individual firing patterns of hippocampal neurons have provided insight
into what is now termed ‘place cells’. Originally discovered in the 1970s, O’Keefe and
Dostrovsky found hippocampal neurons that fired preferentially for certain locations in space.
These cells were promptly named place cells (O’Keefe and Dostrovsky, 1971). Different place
cells were tuned to various fields in space (O’Keefe, 1976). Once established, the ‘place fields’ of
individual cells persisted for many months (Thompson and Best, 1990). This led researchers to
suggest the hippocampus as the brain region responsible for creating an internal map of its
environment and encoding a sense of location (Moser et al., 2015). While these preferentially

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tuned neurons could be shockingly stable, learning readily adapted the firing preference of these
neurons – indicating their ability to remap easily. The preferential firing of these hippocampal
neurons is likely not limited to spatial events, as researchers have found differential firing patterns
during odor-sequence tasks in both CA1 and entorhinal cortex (Ginther et al., 2011; Iragashi et
al., 2014). Other studies have found the existence of ‘time cells’, neurons that encode successive
moments between two events (MacDonald et al., 2011). While the bulk of these studies have
focused on CA3 and CA1 pyramidal neurons, place cells with multiple place fields have been
found in DG as well (Park et al., 2011). Certain place cells fire specifically for borders of the
environment, or the direction of the head (Savelli et al., 2008). Similarly modulated place cells
have been found in the EC as well, indicating this may not be specific to one or two areas within
the hippocampus (Brun et al., 2002). Studies of individual neurons have provided immense
insights into the diversity of functions performed by the hippocampus. Even still, further
knowledge has been gleaned from investigations into individual hippocampal subfields.
1.3 CA1 vs DG
Two of the most well studied hippocampal subfields are CA1 and the DG. The two
regions possess distinct primary excitatory neurons, each displaying different receptor expression,
calcium dynamics, and firing frequencies (Alkadhi, 2019). Pyramidal neurons, the principal
neurons within CA1, are the main output of the hippocampus. They project heavily to the
subiculum and deeper layers of the EC. Some pyramidal neurons also project to areas such as the
amygdala, hypothalamus, PFC, auditory and visual cortex, and NaC (Andersen et al., 1971;
Swanson and Cowan, 1977). The principal neurons of the DG, on the other hand, are dentate

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granule neurons. These excitatory cells receive input from the EC via the perforant path and send
their mossy fiber axons to the pyramidal neurons of CA3 (Amaral et al., 2007).
Pyramidal and granule neurons vary not only in their anatomical projections, but also in
their structural features. Both cell types contain a long apical dendrite. Granule cells have
relatively short and dense dendritic trees, while pyramidal neurons display less compact but more
extensive arborization patterns (Alkadhi, 2019). Pyramidal cell bodies reside in the stratum
pyramidale cell layer. Their apical dendrite extends to the stratum radiatum and arborizes in the
stratum lacunosum moleculare, while their basal dendrites branch in the stratum oriens
(Hammond, 2015). Dentate granule neurons are densely packed in the granule cell layer of the
DG, while their dendrites reside in the molecular layer (Amaral et al., 2007). A typical rat CA1
pyramidal neuron receives around 30k glutamatergic inputs and ~1700 GABAergic inputs
(Megías et al., 2001). Granule neurons, on the other hand, are estimated to receive between 3640-
5600 excitatory inputs – roughly the number of dendritic spines located on their apical dendrites
(Desmond and Levy, 1985; Amaral et al., 2007). Morphological differences help characterize
these two cell types. Nevertheless, it appears that variations within neuronal populations exist as
well.
Increasing evidence points to the principal neurons of both CA1 and DG forming distinct
subpopulations (Fukuda et al., 2003, Amaral et al., 2007; Mizuseki et al., 2011). Pyramidal
neurons differ in their axonal arborization, cell body morphology, dendrite specializations, and
synaptic targets (Mizuseki et al, 2011). In the DG, dendritic tree length varies between dorsal and
ventral granule neurons (Desmond and Levy, 1985). Mature and young granule neurons also
show distinct thresholds for LTP induction (Lopez-Rojas and Kreutz, 2016). It is likely that
differences within and between cell types enhance informational complexity and increase

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diversity of function. For instance, a larger dendritic area may allow for more synaptic inputs –
leading to a greater potential for synaptic plasticity. The plasticity of spines results in rapid
alterations to synaptic firing, allowing synapses to change in response to experience and learning.
Given the same stimuli, CA3-CA1 and perforant path-DG synapses may have entirely
different responses – supporting the notion that these pathways transmit different information
(Huang and Kandel, 2005). For instance, the potassium channel blocker tetraethylammonium
(TEA) induces LTP in CA1 and LTD in DG (Song et al., 2001). Further, fetal alcohol exposure in
rats results in significant neuronal loss in CA1, while DG granule neurons display some
resistance (Miller, 1995; Tran and Kelly, 2003). Variations in output to identical input may be
partially attributed to differences in the density and distribution of postsynaptic receptors. For
instance, granule and pyramidal neuron dendrites express different combinations and levels of
NMDAR subunits (Song et al., 2001; Coultrap et al., 2005). In CA1, there is greater expression of
the GluN2B receptor subunit in comparison to DG (Coultrap et al., 2005). At CA1 hippocampal
synapses, NMDARs provide the primary source of Ca2+ for LTP induction. As such, NMDARs
play a central role in rapid synaptic plasticity and LTP within CA1. Many studies have noted
differences in LTP induction along the dorsoventral axis of CA1 (Schreurs et al., 2017). LTP
appears to have a larger amplitude in dorsal vs ventral CA1. This finding has been corroborated
in the dentate as well, with differences in DG-LTP showing up along the dorsoventral axis
(Schreurs et al., 2017). Variations in the threshold for LTP induction suggest differences in the
type of information encoded.
Variability in the anatomical, structural, and functional properties between neuronal
populations aids the immense complexity of information processing within the hippocampus.
This diversity contributes to learning, memory, and ultimately behavior.

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1.4 Different Molecular Regulators within DG
Uniquely, the dentate gyrus is one of two known brain areas in the human to undergo
adult neurogenesis (Eriksson et al., 1998; Knoth et al., 2010). Many factors affect neurogenesis,
including enriched environment, exercise, chronic stress, aging, and disease (Jessberger et al.,
2009; Schoenfeld and Gould, 2012; Poulose et al., 2017; Babcock et al., 2021; Baptista and
Andrade, 2018). Due to its role in neurogenesis, the DG may be more heavily influenced by
environmental factors such as exercise and learning in contrast to CA1. DG granule neurons also
show more resistance to neuronal death following transient ischemia when compared with CA1
pyramidal neurons, suggesting variability in the molecular mechanisms regulating these neuronal
populations (Kirino, 1982; Pulsinelli, 1985; Krupska et al., 2021).
Mature granule neurons within the dentate gyrus are also unique in that they display a
remarkably low level of intrinsic excitability in comparison with other brain regions (Jung and
McNaughton, 1993; Diamantaki et al., 2016; You et al., 2020). The sparse activation of granule
cells is thought to contribute to pattern separation, the process by which distinct neuronal codes
are assigned to related episodes (Leutgeb et al., 2007; Nakashiba et al., 2012; Berron et al., 2016;
Cayco-Gajic and Silver, 2019). Deficits in pattern separation, as well as overall dysfunction
within the dentate gyrus, are associated with the development of schizophrenia - a debilitating
psychiatric disorder (Das et al., 2014; Yun et al., 2016). Growing clinical and preclinical evidence
points to dentate gyral dysfunction within the prodromal period of schizophrenia, linking this
brain region with progression of the disorder (Tavitian et al., 2019; Nakahara et al., 2020; Jaffe et
al., 2020).

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Differences such as these suggest dentate granule synapses may be regulated by a unique
set of synaptic proteins and molecules. One such molecular regulator is SAP97, a member of the
membrane-associated guanylate kinase (MAGUK) superfamily of proteins. MAGUKs are a
widely expressed protein family integral for the formation and maintenance of synaptic scaffolds
(Oliva et al., 2012). Other members of the MAGUK family include PSD95, PSD93, and SAP102.
While a synaptic role for the latter three proteins has been well documented, a function for SAP97
in synaptic regulation has long remained elusive (Levy et al., 2015; Won et al., 2017). Despite
this, SAP97 is the only MAGUK to directly bind the AMPAR subunit GluA1 (Leonard et al.,
1998; Sans et al., 2001). AMPARs are critical for basal synaptic transmission, as their activation
is fast and transient. They also undergo rapid recruitment during synaptic plasticity (Collingridge
et al., 2009). AMPAR subunits show significant differences in regional expression within the
cortex (Beneyto and Meador-Woodruff, 2004). Molecular quantification of synaptic AMPARs
shows that nearly 80% of synaptic AMPARs in CA1 pyramidal neurons are composed of
GluA1A2 heteromers, whereas the rest are GluA2A3 heteromers (Lu et al., 2009). Subunit
composition of AMPARs has received a great deal of attention, as it has been proposed to dictate
AMPAR trafficking (Bredt and Nicoll, 2003; Shepherd and Huganir, 2007).
Despite SAP97’s direct interaction with GluA1, it was long thought to not share the same
role in synaptic regulation as its DLG subfamily members. One study found that knocking out
SAP97 in Schaffer collateral-CA1 synapses produced no disruptions in synaptic transmission
(Howard et al., 2010). However, our team finds βSAP97 to display robust expression in the
dendrites of DG granule neurons – with significantly lower levels of expression in the pyramidal
neurons of CA1 and CA3 (Kay et al., 2022). βSAP97 knockdown does not alter glutamatergic
neurotransmission in Schaffer collateral-CA1 synapses, while the same genetic manipulation

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produces a drastic increase in synaptic AMPAR function in perforant path-DG granule synapses
(Kay et al., 2022). We find that βSAP97 likely functions at granule synapses by sequestering
GluA1-containing AMPARs perisynaptically (Kay et al., 2022).
Another protein whose synaptic function has long eluded researchers is 4.1N. 4.1N is a
member of the 4.1 family of proteins. The 4.1 family of proteins are multifunctional, membranecytoskeleton adapters which bind transmembrane proteins and link proteins to the plasma
membrane (Baines et al., 2014). Despite binding many synaptic regulatory proteins, including
GluA1, 4.1Ns role in synaptic regulation has remained a mystery (Shen et al., 2000). The only
study to have investigated an in vivo role for 4.1N in synaptic regulation found germline knock
out of 4.1N and 4.1G produced no disruption to glutamatergic synapse function in CA1 pyramidal
neurons (Wozny et al., 2009). Like SAP97, we discover protein 4.1N displays robust dendritic
immunoreactivity in the granule neurons of the DG – with significantly lower levels of expression
in the dendrites of CA3 and CA1 pyramidal neurons. Chapter 2 will detail a novel, postsynaptic
role for protein 4.1N in regulating glutamatergic neurotransmission. Remarkably, within the
hippocampus, this role is specific to DG granule neurons and joins a growing list of synaptic
proteins that play cell-type specific roles in regulating perforant pathway synapses (Roy et al.,
2018; Rao et al., 2019; Grant et al., 2020; Kay et al., 2022). Chapter 3 will highlight a role for
4.1N’s spectrin-actin binding (SAB) domain in augmenting synaptic AMPAR function in DG
granule neurons. This AMPAR phenotype is the opposite of what is seen with loss of 4.1N’s
FERM domain at granule synapses. This work highlights the specificity of 4.1 proteins in
regulating glutamatergic synapses within the hippocampus, while also uncovering that 4.1N is
involved in maintaining stable synaptic AMPAR expression in DG granule neurons in multiple
ways. My hope is that the work in this dissertation contributes to the growing body of literature

11
detailing the molecular specificity between synapse types, and further confirms perforant pathway
synapses are governed by a unique set of molecular regulators.

12
CHAPTER 2: Protein 4.1N Plays a Cell Type-Specific Role in Hippocampal
Glutamatergic Synapse Regulation.
2.1 Abstract
Many glutamatergic synapse proteins contain a 4.1N protein binding domain. However, a role for
4.1N in the regulation of glutamatergic neurotransmission has been controversial. Here, we
observe significantly higher expression of protein 4.1N in granule neurons of the dentate gyrus
(DG granule neurons) compared with other hippocampal regions. We discover that reducing 4.1N
expression in rat DG granule neurons of either sex results in a significant reduction in
glutamatergic synapse function that is caused by a decrease in the number of glutamatergic
synapses. By contrast, we find reduction of 4.1N expression in hippocampal CA1 pyramidal
neurons has no impact on basal glutamatergic neurotransmission. We also find 4.1N’s C-Terminal
Domain to be nonessential to its role in the regulation of glutamatergic synapses of DG granule
neurons. Instead, we show that 4.1N’s FERM domain is essential for supporting synaptic AMPA
receptor function in these neurons. Altogether, this work demonstrates a novel, cell-type specific
role for protein 4.1N in governing glutamatergic synapse function.
2.2 Introduction
Neurons are specialized cells within the central nervous system that transmit information between
each other at synapses. Formation and maintenance of a synapse is dependent on a broad range of
neuropeptides and selectively expressed molecules (Lüscher et al., 2000; Huganir and Nicoll,
2013; Südhof, 2018). Historically, methodological limitations have restricted studies of synaptic
function to one cell type. Results obtained from these isolated investigations are often
extrapolated to deduce whole-brain synaptic regulatory mechanisms. Recent technological
advances have allowed for deeper investigations into the inner workings of the billions of

13
synapses within our brains. Whole-brain synaptome cartography, 3D electron microscopy, and
proteomic mass-spectrometry have uncovered a plethora of brain region-specific synaptic
subtypes, each showing unique morphological and molecular signatures (Roy et al., 2018;
Cizeron et al., 2020; Zhu et al., 2021). While more work remains to be done, it is clear that
population extrapolation undoubtedly masks individual influence when attempting to elucidate
function of an individual protein.
There has been considerable interest in understanding the role protein 4.1N plays in
regulating neuronal function (Shen et al., 2000; Biederer and Südhof, 2001; Wozny et al., 2009;
Lin et al., 2009). 4.1N belongs to a larger 4.1 family of proteins, and is the predominant 4.1
isoform expressed in the brain (Walensky et al., 1999). Biochemical evidence shows protein 4.1N
binds to many glutamatergic synapse regulatory proteins, resulting in 4.1N being deemed a
potential hub protein and central organizer of synaptic membrane proteins (Baines et al.,2014;
Yang et al., 2021). In dissociated hippocampal cultures, 4.1N is enriched along neuronal
dendrites, colocalizing with important glutamatergic synapse proteins (e.g., GluA1 and PSD95)
(Walensky et al., 1999). Protein 4.1N’s ability to interact with neuronal membrane proteins is
made possible through its four-point-one, ezrin, radixin, and moesin (FERM) and C-Terminal
(CTD) domains (Scott et al., 2001), and many of 4.1N’s binding partners show partial or
exclusive postsynaptic protein localization. For example, studies using dissociated rat neurons
show that the AMPA receptor (AMPAR) subunit GluA1 binds to 4.1N (Shen et al., 2000).
Further, 4.1N interacts with SynCAM1, a synaptic cell adhesion molecule that facilitates the
recruitment of AMPARs to glutamatergic synapses (Hoy et al., 2009). However, a role for 4.1N
in synaptic function has been controversial. A germline knockout model of 4.1N and 4.1G
produced no disruptions to basal synaptic transmission or long-term potentiation (LTP) in

14
hippocampal CA1 pyramidal neurons of juvenile mice (Wozny et al., 2009). An independent
study, utilizing an acute lentiviral knockdown of 4.1N in adult mice, found no effect on basal
glutamatergic neurotransmission in CA1 pyramidal neurons - but did observe a reduction in LTP
maintenance (Lin et al., 2009). Based on these studies in CA1 pyramidal neurons, 4.1N’s role in
governing glutamatergic synapse function appears rather modest.
In this study, we investigate whether 4.1N plays a pathway-specific role in regulating
synaptic transmission within the hippocampus. In accordance with previous work, we find that
knockdown of protein 4.1N has no effect on basal glutamatergic neurotransmission in CA1
pyramidal neurons. Our immunohistochemical analysis suggests this lack of a phenotype is due to
very low, if any, protein 4.1N expression in these neurons. Conversely, we observe robust
dendritic protein 4.1N immunoreactivity in granule neurons of the dentate gyrus (DG granule
neurons), and we find that inhibition of protein 4.1N expression in DG granule neurons produces
a marked disruption of both AMPA- and NMDA-receptor mediated neurotransmission. This
disruption of normal synaptic transmission is accompanied by a significant decrease in dendritic
spine density in DG granule neurons. Our molecular replacement experiments demonstrate that
4.1N’s CTD is nonessential to its function at perforant pathway synapses, while 4.1N’s FERM
domain is critical for the observed synaptic AMPAR-mediated deficit. Altogether, our work
demonstrates a novel, cell-type specific function for protein 4.1N in regulating glutamatergic
neurotransmission at perforant pathway synapses and further confirms these synapses are
governed by a unique set of molecular regulators.
2.3 Materials and Methods

15
Experimental Constructs: A previously validated 4.1N-shRNA target sequence against rat 4.1N
was used for all knockdown experiments (5’-AGGAGAGGGATGCGGTATT-3’; Lin et al.,
2009). The 4.1N-shRNA was subcloned behind the H1 promoter region of a GFP-expressing
pFHUGW expression vector. Rat 4.1N cDNA sequence was acquired from Genscript (Clone ID:
ORa13272D). An shRNA-resistant 4.1N was generated by introducing five silent point mutations
within the RNAi target sequence (AGAAGGGGTATGCGATACT; underlined nucleotides
indicate mismatches in the target region). All cloning was performed using overlap-extension
PCR followed by Clontech In-Fusion Cloning (TaKaRa Bio). Both 4.1N cDNA and the shRNAresistant 4.1N cDNA were cloned into a pCAGGS-IRES-mCherry expression vector. The
shRNA-resistant 4.1NΔCTD mutant was generated in house using the method described above,
by deleting the last 144 amino acids corresponding to the CTD domain (Lys1408-Ser1551). The
shRNA-resistant 4.1NΔFERM mutant was generated by deleting the residue corresponding to the
FERM domain (Tyr283 - Leu376) and was obtained from Genscript (Cat. #: SC1626). All
plasmids were confirmed by DNA sequencing. A pFUGW vector expressing only GFP was coexpressed with pCAGGS-IRES-mCherry constructs to enhance identification of transfected
neurons and was also used as a control vector in spine imaging experiments.
Electrophysiology: All experiments were performed in accordance with NIH Guidelines
for the Care and Use of Laboratory Animals, and all procedures were approved by the
Institutional Animal Care and Use Committee of the University of Southern California. 400 μm
rat organotypic entorhino-hippocampal slice cultures were prepared from both male and female
post-natal day 6-8 (P6-8) Sprague Dawley rats as previously described (Stoppini et al., 1991;
Prang et al., 2001; Bonnici and Kapfhammer, 2009; Sadybekov et al., 2017; Tian et al., 2018).
Tissue was isolated and a MX-TS tissue slicer (Siskiyou) was used to make 400 μm transverse

16
sections. Tissue slices were placed on squares of Biopore Membrane Filter Roll (Millipore) and
placed on Millicell Cell Culture inserts (Millipore) in 35 mm dishes. Slices were fed 1mL of
culture media containing MEM + HEPES (Gibco Cat#12360-038), horse serum (25%), HBSS
(25%) and L-glutamine (1 mM). Media was exchanged every other day. Slices with large portions
of entorhinal cortex were visually identified after slicing. These slices were selected and plated
for use in our experiments, and presence of entorhinal cortex was again confirmed when selecting
slices appropriate for data acquisition. Whole-cell recordings were performed on day in vitro 7-8
(DIV7-8). During recordings, slices were maintained in room-temperature artificial cerebrospinal
fluid (aCSF) containing 119 mM NaCl, 2.5 mM KCl, 1 mM NaH2PO4, 26.2 mM NaHCO3 11 mM
glucose, 4 mM CaCl2, and 4 mM MgSO4. 5 μM 2-chloroadenosine and 0.1 mM picrotoxin were
also added to the aCSF to dampen epileptiform activity and block GABAA receptor activity,
respectively. AMPAR-miniature EPSCs (AMPAR-mEPSCs) were isolated by the addition of
0.5μM tetrodotoxin (TTX) and 0.1 mM picrotoxin to the ACSF. Osmolarity was adjusted to 310-
315 mOsm. aCSF was saturated with 95% O2/5% CO2 throughout the recording. Borosilicate
recording electrodes were filled with an internal whole-cell recording solution containing 135
mM CsMeSO4, 8 mM NaCl, 10 mM HEPES, 0.3 mM EGTA, 5 mM QX-314, 4 mM Mg-ATP,
and 0.3 mM Na-GTP. Osmolarity was adjusted to 290–298 mOsm, and pH-buffered at 7.3–7.4.
DG granule neurons and CA1 pyramidal neurons were identified using differential
interference phase contrast microscopy, while GFP-expressing transfected neurons identified
using epifluorescence microscopy. Dual whole-cell recordings of either neuronal subtype were
made through simultaneous recordings from a transfected neuron and a neighboring,
untransfected control neuron. Synaptic responses were evoked by stimulating with a monopolar
glass electrode filled with aCSF in the stratum radiatum for CA1 recordings and the perforant

17
pathway for DG granule neuron recordings. Membrane holding current, pipette series resistance,
and input resistance were monitored throughout recording sessions. Data were acquired using a
Multiclamp 700B amplifier (Molecular Devices), filtered at 2 kHz, and digitized at 10 kHz.
AMPAR-evoked EPSCS (-eEPSCs) were measured at -70mV. NMDAR-eEPSCs were measured
at +40 mV and were temporally isolated by measuring amplitudes 150 ms following the stimulus,
at which point the AMPAR-eEPSC has completely decayed. Paired-pulse ratio was recorded by
delivering two stimuli at varying intervals of 20 ms, 40 ms, 70 ms and 100 ms and dividing the
peak response of stimulus 2 by the peak response of stimulus 1. Data analysis was performed
using Igor Pro (Wavemetrics). In the scatter plots for simultaneous dual whole-cell recordings,
each open circle represents one paired recording, and the closed circle represents the average of
all paired recordings. No more than one paired recording was performed on any given entorhinohippocampal slice.
Biolistic Transfection: Sparse biolistic transfections were performed on DIV1 as
previously described (Stoppini et al., 1991; Schnell et al., 2002; Lu et al., 2009). 50μg of mixed
plasmid DNA was coated on 1μm-diameter gold particles in 0.5 mM spermidine, precipitated
with 0.1 mM CaCl2, and washed four times in pure ethanol. The DNA-coated gold particles were
then coated onto PVC tubing, dried briefly using ultra-pure N2 gas, and stored at 4 °C in
desiccant. Before use, the gold particles were brought up to room temperature and delivered to
slice cultures via a Helios Gene Gun (BioRad). Construct expression was confirmed by GFP or
mCherry epifluorescence.
Immunohistochemistry: P15 Sprague Dawley rats of both sexes were transcardially
perfused with 11 mL of cold PBS and 25 mL of cold 4% PFA in PBS at a flow rate of 3 mL/min.
The hippocampi were immediately dissected and post-fixed overnight at 4 °C in 4% PFA. After 3

18
brief washes in PBS, the hippocampi were sliced using a vibratome at 100μm thickness. Slices
were placed into 24-well culture plates containing PBS and stained within the wells. Slices were
blocked in PBST (PBS + 0.25% TritonX-100) with 10% Goat Serum for 1 h at room temperature,
rinsed in PBST, and incubated with primary antibody diluted in PBST overnight at 4 °C. The next
day, slices were thoroughly washed in PBST and stained with secondary antibody diluted in
PBST for 2 h at room temperature. After this, slices were mounted onto slides, dried for 15 min,
and mounted with either Fluoromount-G (SouthernBiotech Cat#0100-01) or Fluoroshield with
DAPI (Sigma Aldrich Cat#F6057). Slides were imaged and tiled using a Zeiss 880 Confocal
Microscope using a 10x objective. Antibodies used are as follows: monoclonal mouse anti-4.1N
(1:1000, BD Transduction Laboratories Cat#611836, RRID:AB_2098366), goat anti-mouse
Alexa Fluor 488 (1:1000, Thermo Fisher Scientific Cat# A32723, RRID:AB_2633275).
Western blotting: Embryonic day 16.5 (E16.5) rat hippocampi of both sexes were
dissected, dissociated, and cultured in DMEM with 10% FBS. Neurons were plated into 24-well
plates and treated with Lipofectamine™ 2000 (Invitrogen). Plasmid transfections were performed
on DIV2 per manufacturer’s protocol. 25 μL of transfection complex were added to each well.
Lysates were prepared at DIV5 in RIPA buffer containing protease inhibitor mix (Halt™ Protease
Inhibitor Cocktail, EDTA-free (100X), Thermo Fisher Scientific Cat#78425). Proteins were
resolved by SDS-PAGE. Following the transfer, membranes were cut and analyzed by western
blot with a monoclonal mouse anti-4.1N antibody (1:1000, BD Transduction Laboratories
Cat#611836, RRID:AB_2098366) or a polyclonal rabbit anti-GAPDH antibody (1:1000, Thermo
Fisher Scientific Cat#PA1-987, RRID:AB_2107311). Goat anti-Rabbit IgG (H+L)-HRP
Conjugate secondary antibody (1:5000, Bio-Rad Cat# 170-6515, RRID:AB_11125142) or goat
anti-Mouse IgG (H+L)-HRP Conjugate secondary antibody (1:5000, Thermo Fisher Scientific

19
Cat# 31430, RRID:AB_228307) were used for all immunoblotting experiments described. All
lysates were run on a 4 –15% Mini-PROTEAN TGX Precast Protein Gel (Bio-Rad) with 50 μg of
protein loaded per lane. Membranes were scanned using the Bio-Rad Chemidoc Imaging System.
Spine Density Analysis: Cultured entorhino-hippocampal slices were transfected on DIV1
with pFUGW-GFP construct, pFHUGW-GFP-shRNA construct, or pFHUGW-GFP-shRNA +
pCAGGS-mCherry-cDNA constructs. Slices were fixed in 4% PFA, 4% sucrose in PBS, and
washed three times with PBS, then cleared with an abbreviated SeeDB-based protocol (Ke et al.,
2013) and mounted on microscope slides. Images were acquired at DIV7 using super-resolution
microscopy (Carl Zeiss). High-resolution confocal z-stacks of spine-containing DG granule
neuron secondary apical dendrites were acquired on a Zeiss 880 using an EC Plan-Neofluar
40x/1.3 oil-immersion DIC M27 objective. ~60 μm sections of secondary apical dendrites were
manually selected for analysis. Z-stacks were collected at maximum X–Y pixel dimensions (512
x 512 pixels) at 8 bits with a 488 nm laser excitation wavelength. An experimenter, blinded to the
experimental condition, performed spine density analysis on sections using the Dendritic Spine
Counter plug-in on ImageJ to count spines extending laterally from the dendrite. The ImageJ
plug-In SpineJ was used to obtain values for the following spine morphology metrics: spine head
area, total spine length, neck width, and head width (used for spine type classification) (Levet et
al., 2020). Dendritic processes are commonly classified into the following categories: stubby,
mushroom, and thin. Based on previous literature, we categorized dendrites into three
aforementioned categories using the following criteria. Stubby spines are those which lack a
visible neck region but have a bulbous head. Mushroom spines, often viewed as mature spines,
are those with a short length (<2um), a clearly defined neck region, and a head which is >50% as

20
wide as the neck. Thin spines are those with a long length (>2um) as well as a head (Mattison et
al., 2014; Lin et at., 2004).
Experimental Design and Statistical Analyses: All electrophysiological data are expressed
as mean ± standard error measurement (SEM). Imaging analysis was performed blind to
experimental condition. Statistical significance was determined using: Wilcoxon signed-rank test
for paired dual whole-cell patch clamp data, Wilcoxon rank-sum test for electrophysiological data
across independent conditions as well as imaging data, and Student’s t test for paired-pulse
facilitation data. Data were analyzed using IGOR Pro (Wavemetrics RRID:SCR_000325) or
KaleidaGraph (Synergy Software RRID:SCR_014980), and graphed using Microsoft Excel
(RRID:SCR_016137) or GraphPad Prism (RRID:SCR_002798). All p-values <0.05 were
considered significant and denoted with a single asterisk. All error bars represent standard error
measurement. For all experiments, at least 4 male and female rat pups were used. Sample sizes in
the present study are similar to those reported in the literature (Herring and Nicoll, 2016; Incontro
et al., 2018).
Coefficient of variation (CV) analysis was performed on AMPAR-eEPSCs by comparing
the change in eEPSC variance with the change in mean amplitude as previously described (Del
Castillo and Katz, 1954; Bekkers and Stevens, 1990; Malinow and Tsien, 1990; Xiang et al.,
1994). CV was calculated as SD/M (SD = standard deviation; M = mean). The SD and M were
measured, normalized and plotted for a concurrent set of stimuli from a control and its
neighboring, transfected cell. Theoretical and experimental work shows that CV−2 (M2
/SD2
) is
invariant with changes in quantal size (i.e., the number of AMPARs at all synapses), while
CV−2 varies predictably with changes in quantal content (i.e., the number of functional synapses
containing AMPARs) according to the following equation: CV−2 = n × Pr/(1 − Pr). In this

21
equation, n is the number of vesicle release sites and Pr is the presynaptic release probability.
CV−2 values for transfected and control cells were plotted on the y-axis, and mean eEPSC
amplitude values for transfected and control cells were plotted on the x-axis. Values on or near
the 45° (y = x) line indicate changes in quantal content, while values approaching the horizontal
line (y = 1) indicate a change in quantal size. Filled circles represent the mean ± SEM of the entire
dataset.
Failure analysis was performed by analyzing AMPAR-eEPSCs from dual whole-cell
patch-clamp recordings where stimulation levels elicited failures visually distinguishable from
synaptic currents. Stimulation events were assigned as failures if their absolute magnitudes were
less than or equal to noise for each sweep. The number of failures for each cell was estimated as
the number of stimulation events with absolute current amplitude not greater than noise divided
by the total number of stimulation events to yield the percentage failure rate.
Data, Materials, and Software Availability: All study data are provided within the paper.
All additional information will be made available upon reasonable request to the authors.
2.4 Results
Protein 4.1N is required for the structure and function of glutamatergic synapses in DG granule
neurons
Previous work has shown that knocking out protein 4.1N expression in CA1 pyramidal neurons of
the hippocampus does not affect basal glutamatergic synapse function (Wozny et al., 2009; Lin et
al., 2009). However, gene expression profiles of different hippocampal subregions show
extensive differences in RNA and synaptic protein expression (Datson et al., 2009; Rao et al.,
2019; Kay et al., 2022). To determine the expression profile of protein 4.1N, we performed

22
immunohistochemical analysis on hippocampal slices from rats of either sex. This analysis
revealed a marked contrast between the expression of protein 4.1N in the DG compared to other
hippocampal subregions. The inner and outer molecular layer of the DG display robust
expression, whereas the stratum oriens and stratum radiatum of CA3 and CA1 exhibit little, if
any, 4.1N expression (Fig. 1A).
Given the DG granule neuron-specific 4.1N expression we observe in the hippocampus,
we sought to determine whether protein 4.1N has a functional role at DG granule neuron
synapses. To accomplish this, we generated a 4.1N-shRNA construct utilizing a previously
published shRNA sequence against 4.1N (Lin et al., 2009). We then independently verified the
validity of our knockdown in dissociated hippocampal neurons using a well-established antibody
against protein 4.1N (Lin et al., 2009) (Fig. 1B). Western blot analysis of hippocampal neuron
homogenates revealed that expression of this 4.1N-shRNA reduces 4.1N expression by ~70%.
Following this, we employed a biolistic transfection method to express our protein 4.1N-shRNA
in neurons in rat organotypic entorhino-hippocampal slice cultures (Elias et al., 2008; Paskus et
al., 2019; Tian et al., 2021). These slices allow us to examine the impact genetic modifications
have on hippocampal neurons within their native circuitry (Stoppini et al., 1991; Schnell et al.,
2002). 6 days after transfection, we record AMPA- and NMDA-receptor evoked excitatory
postsynaptic currents (AMPAR- and NMDAR-eEPSCs) from transfected and neighboring,
untransfected control DG granule neurons simultaneously during perforant pathway stimulation
(Fig. 1C). This approach allows for a pairwise, internally controlled comparison of the
consequences of our acute genetic manipulation. We find that knockdown of protein 4.1N in DG
granule neurons produces a ~60% reduction in both AMPAR-eEPSC amplitude (n = 11 pairs, p =
0.032, Wilcoxon signed-rank test; Fig. 1D) and NMDAR-eEPSC amplitude (n = 10 pairs, p =

23
0.0019, Wilcoxon signed-rank test; Fig. 1E). To determine whether this reduction is due to a
disruption in presynaptic neurotransmitter release, we performed paired-pulse facilitation (PPF)
experiments in DG granule neurons. We observe no changes in PPF at a range of interstimulus
intervals (ISIs) between neurons transfected with a 4.1N-shRNA when compared to
untransfected, control neurons (n = 6 pairs, ISI: 20ms: p = 0.62, 40ms: p = 0.90, 70ms: p = 0.47,
100ms: p = 0.50, Student’s t test; Fig. 1F). Thus, the observed deficits in glutamatergic synapse
function seen with knockdown of 4.1N are due to a reduction in postsynaptic function.
Most excitatory synapses are formed on dendritic spines. Observed reductions in both
AMPAR- and NMDAR-eEPSC amplitude are often associated with a reduction in dendritic spine
density. Therefore, we performed dendritic spine density analysis to determine whether the
observed deficits in AMPA- and NMDA-receptor mediated transmission caused by 4.1N
knockdown are produced by a loss of spines. We compared the density of dendritic spines on
secondary apical dendrites of DG granule neurons expressing the 4.1N-shRNA or a control GFP
construct (GFP: n =31 segments, 4.1N-shRNA: n =37 segments, p< 0.0001, Wilcoxon rank-sum
test; Fig. 1G). We find that knocking down protein 4.1N produces a ~50% reduction in dendritic
spine density in DG granule neurons. The magnitude of this reduction in spine density is
comparable to the percent reduction we see in AMPAR- and NMDAR-eEPSC amplitude
following knockdown of protein 4.1N expression (Fig. 1D, E). We also examined the total spine
length, head area, and proportion of different spine types (mushroom, thin, stubby) in 4.1NshRNA-expressing neurons compared to GFP-expressing control neurons, and found no
significant differences in these spine parameters in 4.1N-shRNA-expressing DG granule neurons
(GFP: n = 5 cells, 4.1N-shRNA: n = 4 cells, p = 0.90 for spine length, p = 1 for head area, p =
0.56 for stubby spines, p = 0.90 for thin spines, p = 0.29 for mushroom spines, Wilcoxon rank-

24
sum test; Fig 1G). Taken together, these data demonstrate that 4.1N is essential for maintaining
both the structure and function of glutamatergic synapses in DG granule neurons.
We also examined whether knockdown of 4.1N alters single synaptic events in DG
granule neurons by recording AMPAR-mediated miniature excitatory postsynaptic currents
(AMPAR-mEPSCs). Consistent with the reduction in dendritic spine number we observe, we find
that the frequency of AMPAR-mEPSCs is reduced by ~75% in 4.1N knockdown neurons
(control: n = 6 cells, 4.1N-shRNA: n = 6 cells, control = 0.62 ± 0.11 Hz, 4.1N-shRNA = 0.14 ±
0.02 Hz, p = 0.002, Wilcoxon rank-sum test; Fig. 1H). The amplitude of the remaining AMPARmEPSCs in 4.1N-shRNA-expressing neurons was unchanged (control: n = 6 cells, 4.1N-shRNA:
n = 6 cells, control = 14.33 ± 1.12 pA; 4.1N-shRNA = 20.01 ± 2.38 pA, p = 0.13, Wilcoxon ranksum test; Fig. 1H). These data provide additional confirmation that reducing 4.1N protein
expression results in a substantial loss of functional glutamatergic synapses in DG granule
neurons.
It has been shown previously that 4.1N preferentially binds to the AMPAR subunit
GluA1, but not to GluA2 or GluA3 (Coleman et al., 2003). Previous work has also shown that
decay kinetics of synaptic AMPAR currents vary by subunit composition, with GluA2/A3
heteromers deactivating more quickly than GluA1/A2 heteromers (Lu et al., 2009; Herring et al.,
2013). Therefore, the loss of GluA1-containing AMPARs at synapses produces a hallmark
increase in the decay rate of synaptic AMPAR currents caused by a higher GluA2/A3 to
GluA1/A2 synaptic AMPAR ratio (Lu et al., 2009; Herring et al., 2013). To determine whether
the synaptic AMPAR deficit seen with loss of 4.1N is accompanied by a change in synaptic
AMPAR subunit composition, we examined the kinetics of AMPAR-eEPSCs. We find no
differences in the AMPAR-eEPSC rate of decay between 4.1N-shRNA-expressing and control

25
DG granule neurons (n = 9 pairs, p = 0.73, Wilcoxon signed-rank test; Fig. 1I). Altogether, these
data demonstrate that reducing 4.1N expression in DG granule neurons results in a substantial
reduction in glutamatergic synapse number in DG granule neurons.
4.1N knockdown does not perturb basal glutamatergic synapse function in CA1 pyramidal
neurons
Our immunohistochemical analysis reveals considerably lower expression of protein 4.1N in the
dendrites of CA3 and CA1 pyramidal neurons when compared to the dendrites of DG granule
neurons (Fig. 2A). Such data suggest that the synaptic expression of protein 4.1N is very low in
hippocampal subregions outside of the DG, and therefore may play a DG-specific role in synaptic
regulation within the hippocampus. To test this hypothesis, we examined the glutamatergic
synapse function of CA1 pyramidal neurons transfected with our 4.1N-shRNA construct
compared to that of neighboring, untransfected control neurons (Fig. 2B). Consistent with our
immunohistochemical data, we observe no significant change in either AMPAR- (n = 8 pairs, p =
0.55, Wilcoxon signed-rank test; Fig. 2C) or NMDAR-eEPSC amplitude in CA1 pyramidal
neurons (n = 8 pairs, p = 0.95, Wilcoxon signed-rank test; Fig. 2D). We also examined whether
presynaptic neurotransmitter release is altered in 4.1N knockdown neurons. We observe no
change in PPF at a range of paired stimulus intervals between CA1 pyramidal neurons transfected
with our 4.1N-shRNA when compared to untransfected, control neurons (n = 6 pairs, ISI: 20ms: p
= 0.91, 40ms: p = 0.41, 70ms: p = 0.32, 100ms: p = 0.48, Student’s t test; Fig. 2E). Due to 4.1N’s
proposed interaction with the AMPAR subunit GluA1, we again examined AMPAR-eEPSC
decay rate in CA1 pyramidal neurons (Shen et al., 2000; Coleman et al., 2003). We find the decay
rates of AMPAR-eEPSCs from 4.1N-shRNA-expressing neurons to not vary from that of control

26
neurons (n = 7 pairs, p = 0.94, Wilcoxon signed-rank test; Fig. 2F). Akin to other studies, we
conclude that 4.1N has no role in basal synaptic transmission in CA1 pyramidal neurons (Wozny
et al., 2009; Lin et al., 2009). Together, these data demonstrate that 4.1N plays a cell-type specific
role in regulating glutamatergic neurotransmission in the hippocampus.
Protein 4.1N’s C-Terminal Domain is not critical to its function at DG granule synapses
To further confirm the specificity of our protein 4.1N-shRNA, we generated a recombinant
shRNA-resistant 4.1N cDNA expression construct. We then co-expressed the 4.1N-shRNA with
the shRNA-resistant 4.1N cDNA (4.1N Rescue) in DG granule neurons and performed paired
recordings. We find that molecularly replacing endogenous 4.1N with recombinant 4.1N rescues
the reductions in AMPAR- (n = 9 pairs, p = 1, Wilcoxon signed-rank test; Fig. 3B, D) and
NMDAR-eEPSC amplitude caused by expression of 4.1N-shRNA (n = 9 pairs, p = 0.57,
Wilcoxon signed-rank test; Fig. 3E, G). We also examined spine density phenotypes of the 4.1N
rescue construct in comparison to 4.1N-shRNA-expressing and GFP-expressing control DG
granule neurons. Co-expression of 4.1N-shRNA with 4.1N-shRNA-resistant cDNA rescues the
reduction in dendritic spine density produced by our 4.1N-shRNA (GFP: n =31 segments, 4.1NshRNA & 4.1N-shRNA-resistant cDNA (4.1N Rescue): n = 48 segments, p = 0.27, Wilcoxon
rank-sum test; Fig. 3I). Further, we find no differences in any measures of spine morphology
between 4.1N Rescue-expressing DG granule neurons and control, GFP-expressing DG granule
neurons (GFP: n = 5 cells, 4.1N Rescue: n = 6 cells, p = 0.66 for spine length, p = 0.54 for head
area, p = 0.41 for stubby spines, p = 0.66 for thin spines, p = 1 for mushroom spines, Wilcoxon
rank-sum test; Fig. 3I). We, therefore, conclude that the deficit in glutamatergic synapse density

27
and function we observe resulting from expression of our 4.1N-shRNA is due to reduced protein
4.1N expression.
Having confirmed that the observed synaptic deficit seen in DG granule neurons following
loss of 4.1N is rescued by recombinant 4.1N, we sought to understand the 4.1N domain(s)
responsible for producing these marked decreases in AMPA- and NMDA-receptor mediated
currents. Initially, we hypothesized that 4.1N function at DG granule synapses was dependent on
its C-Terminal domain (CTD), given that this domain was previously reported to interact with the
AMPA receptor subunit GluA1 (Shen et al., 2000; Coleman et al., 2003). Based on this reported
interaction, we hypothesized that molecular replacement of endogenous 4.1N with a truncated
variant missing its CTD (4.1N∆CTD) would fail to rescue synaptic deficits seen with knockdown
of full length 4.1N (Fig. 3A). Surprisingly, molecular replacement of endogenous 4.1N with
4.1N∆CTD rescues both AMPAR- (n = 7 pairs, p = 0.94, Wilcoxon signed-rank test; Fig. 3C, D)
and NMDAR-eEPSC amplitudes (n = 7 pairs, p = 0.69, Wilcoxon signed-rank test; Fig. 3F, G) in
DG granule neurons. Given this unexpected finding, we sought to examine whether this rescue of
synaptic function results from a compensatory effect of remaining GluA2/A3 subunits. As
previously mentioned, GluA2/A3 heteromers deactivate at faster rates than GluA1/A2 heteromers
(Lu et al., 2009; Herring et al., 2013). However, we find the kinetics of AMPAR-eEPSCs from
4.1N∆CTD-expressing neurons to not vary from that of control neurons (n = 6 pairs, p = 0.56,
Wilcoxon signed-rank test; Fig. 3H). These data strongly suggest that the cell-type specific
synaptic deficits in DG granule neurons following 4.1N knockdown are not due to a loss of
protein-protein interactions supported by 4.1N’s CTD.

28
Protein 4.1N’s FERM domain is required for synaptic AMPA receptor function in DG granule
neurons.
4.1N’s remaining protein binding domain, the FERM domain, binds a vast array of
transmembrane proteins (Biederer and Südhof, 2001; Scott et al., 2001; Hoy et al., 2009; Li et al.,
2007). Given that elimination of 4.1N’s CTD does not affect 4.1N function, we hypothesized that
the FERM domain may play an important role in supporting protein 4.1N’s function at
glutamatergic synapses in DG granule neurons. To determine the importance of protein 4.1N’s
FERM domain, we molecularly replaced endogenous 4.1N with a mutant variant lacking its
FERM domain (4.1NΔFERM; Fig. 4A). We find that replacing endogenous 4.1N with
4.1NΔFERM fails to rescue the reduction in AMPAR-eEPSC amplitude caused by 4.1N
knockdown (n = 13 pairs, p = 0.021, Wilcoxon signed-rank test; Fig. 4B, C). However,
4.1NΔFERM is able to rescue the reduction in NMDAR-eEPSC amplitude caused by 4.1N
knockdown (n = 12 pairs, p = 0.52, Wilcoxon signed-rank test; Fig. 4D, E). As such, we
examined whether removal of 4.1N’s FERM domain is accompanied by a change in the number
and/or morphology of dendritic spines at DG granule synapses. We find that co-expression of
4.1N-shRNA with 4.1NΔFERM cDNA rescues the reduction in spine density produced by
reducing protein 4.1N expression (GFP: n =31 segments, 4.1N∆FERM: n = 31 segments, p= 0.59,
Wilcoxon rank-sum test; Fig. 4F). Further, we find no significant differences in any spine
parameters assessed in DG granule neurons co-expressing 4.1N-shRNA and 4.1N∆FERM
compared to GFP-expressing control neurons (GFP: n = 5 cells, 4.1N∆FERM: n = 6 cells, p =
0.79 for spine length, p = 0.43 for head area, p = 0.36 for stubby spines, p = 0.66 for thin spines, p
= 0.93 for mushroom spines, Wilcoxon rank-sum test; Fig. 4F). Next, we examined the frequency
and amplitude of AMPAR-mEPSCs to assess whether removal of the FERM domain produces an

29
alteration in single synaptic events. We find that AMPAR-mEPSC frequency is reduced by ~65%
in 4.1N∆FERM mutants (control: n = 6 cells, 4.1N∆FERM: n = 6 cells, control = 1.31 ± 0.28 Hz,
4.1N∆FERM = 0.45 ± 0.09 Hz, p = 0.041, Wilcoxon rank-sum test; Fig. 4G), while the amplitude
of remaining AMPAR-mEPSCs is comparable to that of control DG granule neurons (control: n =
6 cells, 4.1N∆FERM: n = 6 cells, control = 14.28 ± 1.67 pA, 4.1N∆FERM = 19.17 ± 3.54 pA; p =
0.39, Wilcoxon rank-sum test; Fig. 4G). Together, these data demonstrate that protein 4.1N’s
FERM domain plays a specialized role in synaptic AMPAR function in DG granule neurons.
Loss of 4.1N’s FERM domain results in a selective deficit in AMPAR, but not NMDAR,
function. Loss of 4.1N’s FERM domain also results in a reduction in AMPAR-mEPSC frequency,
but AMPAR-mEPSC amplitude is unchanged. Such data are consistent with the deletion of
4.1N’s FERM domain causing a loss of all AMPARs at a subset of existing synapses, resulting in
an increase in the number of “silent synapses”. To further examine this possibility, we performed
coefficient of variation (CV-2
) analysis on AMPAR-eEPSC amplitudes (Del Castillo and Kats
1954, Malinow and Tsien 1990). By comparing the normalized variance in AMPAR-eEPSC
amplitudes from control and transfected neurons receiving the same stimulus, we are able to
estimate the relative quantal size and quantal content. Changes in quantal size modify the mean
eEPSC amplitude and variance such that the CV-2 remains constant, and indicate a change in the
number of glutamate receptors at all synapses. In contrast, changes in quantal content produce
proportional changes of equal magnitude in CV-2 and mean eEPSC amplitudes which cause the
marker of the mean to fall on the diagonal (y=x) line. Changes in quantal content indicate a
change in the number of synapses expressing glutamatergic receptors. CV-2 analysis reveals that
the reduction in AMPAR-eEPSC amplitude caused by loss of the FERM domain is due to a
reduction in quantal content and, therefore, due to an increase in the number of silent synapses in

30
these neurons (n = 13 pairs, Fig. 4H). An increase in silent synapse number is also accompanied
by an increase in the probability that a given axonal stimulation fails to elicit an AMPAR-eEPSC
(Goold and Nicoll, 2010; Gray et al., 2011). We, therefore, examined the frequency at which
presynaptic stimulation fails to elicit an AMPAR-eEPSC when 4.1N’s FERM domain is deleted.
Consistent with our AMPAR-mEPSC data and CV-2 analysis, we find that removal of 4.1N’s
FERM domain results in a significant increase in failures (n = 13 pairs, p = 0.035, Student’s t test;
Fig. 4I). Altogether, our data demonstrate that 4.1N’s FERM domain is critical for synaptic
AMPAR expression and that loss of 4.1N’s FERM domain increases the percentage of synapses
that lack functional AMPARs.
It has been shown previously that loss of GluA1-containing AMPARs results in a
significant reduction in AMPAR-eEPSC amplitude, no change in NMDAR-eEPSC amplitude,
and a substantial reduction in AMPAR-mEPSC frequency (Lu et al., 2009; Herring et al., 2013).
Removal of 4.1N’s FERM domain produces a similar synaptic phenotype, and 4.1N has been
proposed to support the trafficking of GluA1-containing AMPARs (Lin et al., 2009; Bonnet et al.,
2023). Given that the loss of GluA1-containing AMPARs also produces a hallmark speeding of
synaptic AMPAR current decay kinetics (Lu et al., 2009; Herring et al., 2013), we examined
AMPAR-eEPSC decay kinetics in the context of a selective deletion of 4.1N’s FERM domain.
Indeed, we find that removal of 4.1N’s FERM domain produces a marked speeding in the decay
of AMPAR-eEPSCs (n = 9 pairs, p = 0.039, Wilcoxon signed-rank test; Fig. 4J). Thus, the loss of
4.1N’s FERM domain phenocopies the loss of the GluA1 subunit in neurons and strongly
suggests that 4.1N’s FERM domain, rather than its CTD, plays a critical role in the expression of
functional GluA1-containing AMPARs in a subpopulation of DG granule neuron synapses.

31
2.5 Discussion
We have identified, for the first time, a postsynaptic role for protein 4.1N in regulating basal
glutamatergic neurotransmission. Within the hippocampus, this function is cell-type specific.
Consistent with previous work, we show that knockdown of protein 4.1N in CA1 pyramidal
neurons has no effect on basal glutamatergic neurotransmission (Wozny et al., 2009; Lin et al.,
2009). By contrast, we find that reducing protein 4.1N expression in DG granule neurons leads to
a substantial reduction in basal glutamatergic synapse function. Immunohistochemical analysis
supports the validity of these results, showcasing 4.1N protein expression levels to be markedly
greater in DG granule neurons in comparison to the pyramidal neurons of CA1 and CA3.
Here we show that knockdown of protein 4.1N in DG granule neurons results in a loss of
glutamatergic synapses. Reduced protein 4.1N expression in DG granule neurons produces
significant reductions in both AMPAR- and NMDAR- eEPSC amplitude. These deficits in
synaptic function are accompanied by a comparable reduction in dendritic spine density. Spine
morphology analysis and AMPAR-mEPSC data suggest that those synapses which remain
following 4.1N knockdown are normal, and are potentially supported by residual endogenous
4.1N expression. Both the structural and functional synaptic deficits produced by reducing protein
4.1N expression in DG granule neurons are rescued with the expression of recombinant protein
4.1N. Based on these data, we conclude that protein 4.1N plays a major and cell-type specific role
in supporting glutamatergic synapse structure and function in DG granule neurons of the
hippocampus.
Having identified a major role for 4.1N in DG granule neuron glutamatergic synapse
regulation, we examined which domains of protein 4.1N support glutamatergic synapse function
in these neurons. Protein 4.1N contains two established domains capable of protein-protein

32
interactions: the C-Terminal domain (CTD) and the FERM domain. Initially, we hypothesized
that 4.1N’s postsynaptic function at glutamatergic synapses in DG granule neurons was
dependent on its CTD. Previous biochemical studies have reported that 4.1N’s CTD binds to a
number of glutamate receptor subunits, including the AMPAR subunit, GluA1 (Shen et al., 2000;
Coleman et al., 2003). In dissociated neuronal cultures, 4.1N has been implicated in synaptic
GluA1 trafficking, suggesting a noteworthy role for 4.1N in AMPAR regulation (Lin et al., 2009,
Bonnet et al., 2023). However, the 4.1N domain required for this role was not examined.
Surprisingly, we find that expression of recombinant 4.1N lacking its CTD in DG granule
neurons completely rescues the 4.1N knockdown phenotype. Based on these data, we conclude
that 4.1N’s CTD is not necessary for the role protein 4.1N plays in supporting synaptic
transmission at perforant pathway synapses.
Having found the CTD to be dispensable for 4.1N’s synaptic regulatory role in DG
granule neurons, we examined the role of the FERM domain. This domain has been shown to
bind to a large number of postsynaptic regulatory proteins implicated in the assembly,
maintenance, and plasticity of synapses (Biederer and Südhof, 2001; Hoy et al., 2009; Li et al.,
2007; Cohen et al., 1998). Molecular replacement of endogenous 4.1N with a mutant lacking the
FERM domain (4.1N∆FERM) only partially restores synaptic deficits seen with knockdown of
4.1N. While NMDAR function and dendritic spine density are restored to baseline, synaptic
AMPAR function remains markedly reduced with the 4.1N∆FERM mutant. Our AMPARmEPSC, quantal analysis, and failure analysis data reveal that the selective reduction in AMPAReEPSC amplitude is due to a reduction in the number of glutamatergic synapses that express
functional AMPARs. Together, these data demonstrate that 4.1N’s FERM domain is critical for

33
supporting synaptic AMPAR expression in DG granule neurons and that its dysfunction results in
an increase in the number of silent synapses.
As stated, protein 4.1N has been implicated in the trafficking of GluA1-containing
AMPARs (Lin et al., 2009; Bonnet et al., 2023). It has been shown that knocking out GluA1 and
preventing the trafficking of GluA1-containing receptors results in a marked reduction in
AMPAR-eEPSC amplitude and AMPAR-mEPSC frequency (Lu et al., 2009; Herring et al.,
2013). We find that deletion of 4.1N’s FERM domain produces a similar synaptic phenotype.
Additionally, it has been shown that preventing GluA1 expression and trafficking results in a
marked speeding of synaptic AMPAR current decay kinetics. This effect is attributable to the loss
of GluA1/A2 receptors, which deactivate more slowly than GluA2/A3 receptors (Lu et al., 2009;
Herring et al., 2013). We, therefore, examined the impact of 4.1N’s FERM domain on AMPAReEPSC decay. Remarkably, we find that deletion of 4.1N’s FERM domain produces a significant
increase in the decay rate of AMPAR-eEPSCs - similar to that observed with the loss of GluA1
expression/trafficking. Thus, loss of 4.1N’s FERM domain phenocopies the synaptic alterations
observed when GluA1-containing AMPARs are compromised in neurons (Lu et al., 2009;
Herring et al., 2013). Together, these results suggest that, in DG granule neurons, 4.1N’s FERM
domain, as opposed to its CTD, is responsible for GluA1-containing AMPAR trafficking to
synapses.
In the present study, we show that knocking down 4.1N expression in DG granule neurons
results in a reduction in AMPAR-and NMDAR-eEPSC amplitudes that is caused by a loss of
dendritic spines. Molecular replacement of 4.1N with 4.1N∆FERM rescues dendritic spine loss
and supports synapses that contain NMDARs but lack AMPARs. Thus, 4.1N’s FERM domain is
necessary for the presence of functional AMPARs at synapses. However, the domain(s) required

34
for 4.1N’s role in supporting glutamatergic synapse structure remains unknown. We hypothesize
that a yet to be identified domain of 4.1N promotes the recruitment of actin regulatory proteins
that are required for the development or maintenance of dendritic spines. Alternatively, it is
possible that 4.1N directly supports dendritic NMDAR recruitment and/or clustering - which in
turn facilitates NMDAR-dependent synaptogenesis (Kwon and Sabatini, 2011). At present, a
direct interaction between 4.1N and NMDARs has yet to be observed. Nevertheless, it is plausible
that 4.1N promotes dendritic NMDAR recruitment/clustering through an intermediate protein –
and that the loss of this indirect interaction following 4.1N knockdown inhibits NMDARmediated synaptogenesis. Protein 4.1 is often described as a hub protein with the ability to
associate and organize many synaptic regulatory proteins (Baines et al.,2014; Yang et al., 2021).
Going forward, it will be important to carry out thorough investigations of candidate binding
proteins in order to identify the protein-protein interactions necessary for the role 4.1N plays in
supporting the structure and function of glutamatergic synapses in DG granule neurons.
In the present work, we do not observe a role for 4.1N in supporting basal glutamatergic
synapse function in CA1 pyramidal neurons. Historically, a synaptic regulatory role for 4.1N in
CA1 pyramidal neurons has been controversial. Germline knockout of 4.1N and 4.1G, another 4.1
family member, in a juvenile mouse model produced no changes in baseline synaptic transmission
or in LTP in CA1 pyramidal neurons (Wozny et al., 2009). In an independent study, lentiviral
knockdown of 4.1N in CA1 pyramidal neurons of adult mice resulted in no change to basal
glutamatergic synapse function, but LTP maintenance was found to be inhibited (Lin et al., 2009).
These seemingly discrepant results may suggest that 4.1N has a developmentally regulated role in
LTP maintenance in CA1 pyramidal neurons. In our study we observe little, if any, 4.1N
expression in the CA1 hippocampal subregion of juvenile rats. However, protein 4.1N expression

35
within CA1 may achieve appreciable levels later in adulthood. Going forward, it will be important
to examine cell-type specific roles for 4.1N in synaptic function and plasticity at different stages
of development.
In conclusion, we demonstrate a major role for 4.1N in supporting basal glutamatergic
synapse structure and function, and we find that this role is cell-type specific within the
hippocampus. In contrast to a possible modulatory role in LTP maintenance observed in CA1
pyramidal neurons (Lin et al., 2009), we believe that 4.1N serves as a master regulator of synaptic
development in DG granule neurons. With this study, 4.1N joins a growing list of synaptic
proteins that play cell-type specific roles in regulating perforant pathway synapses (Roy et al.,
2018; Rao et al., 2019; Grant et al., 2020; Kay et al., 2022). For example, we have recently shown
that schizophrenia-associated mutations in the synaptic scaffolding protein SAP97 produce
dramatic and likely pathological elevations in glutamatergic synapse strength specifically in DG
granule neurons (Kay et al., 2022). Glutamatergic synapses onto DG granule neurons act as the
gateway for information flow into the hippocampus. Regulation of the number and strength of
these synapses underlie the ability of the dentate gyrus to separate similar information into
distinct representations in a process called pattern separation (Leutgeb et al., 2007; McHugh et al.,
2007). It is generally held that pattern separation underlies our ability to distinguish a memory
from similar stored memories. Thus, functional deficits in proteins governing dentate gyral
synaptic function stand to have profound consequences on how we perceive the external world.
Going forward, it will be important to understand how the unique synaptic proteins present at
perforant pathway synapses support information processing in the dentate gyrus, and how deficits
in the function of these proteins contribute to the development of complex brain disorders.

36
Figure 2.1 Protein 4.1N is required for glutamatergic synapse structure and function in DG
granule neurons.
(A) Representative immunolabeling showing enrichment of protein 4.1N in the molecular layer of
the DG in a rat hippocampal slice. Blue box shows enlarged DG region. GL: granule layer, ML:

37
molecular layer. (B) Western blot showing shRNA-mediated reduction of 4.1N protein in
dissociated hippocampal neurons. (C) Schematic representation of electrophysiological recording
setup for DG granule neurons. (D) Knocking down 4.1N significantly decreases both AMPAReEPSC (n = 11 pairs) and (E) NMDAR-eEPSC amplitudes (n = 10 pairs) in DG granule neurons.
(D-E) Scatterplots show eEPSC amplitudes for pairs of untransfected and transfected cells (open
circles) with corresponding mean ±SEM (filled circles). Insets show representative current traces
from control and transfected (blue) neurons with stimulation artifacts removed. Scale bars: 20 ms,
20pA for both AMPAR-eEPSCs and NMDAR-eEPSCs. Bar graphs show the average AMPAReEPSC and NMDAR-eEPSC amplitudes (±SEM) of DG granule neurons expressing the 4.1NshRNA (blue) normalized to their respective control cell average eEPSC amplitudes (black). (F)
Paired-pulse facilitation ratios (mean ±SEM) for 4.1N-shRNA-expressing DG granule neurons
and paired control neurons show no detectable differences in facilitation at a variety of
interstimulus intervals (ISIs) (n = 6 pairs, ISI: 20, 40, 70, and 100 ms). Peak 1-scaled current
traces from control (black) and transfected (blue) neurons. Scale bars: 20 ms. (G) 4.1N-shRNAexpressing DG granule neurons have significantly lower dendritic spine density in comparison to
GFP-expressing control neurons. Leftmost images display representative dendritic segments of
GFP- (left) and 4.1N-shRNA- (right) expressing DG granule neurons. Scale bars: 10µm. Violin
plots show a significant difference in the spine density of DG granule neurons expressing the
4.1N-shRNA when compared to GFP-expressing control neurons (GFP: n =31 segments, 4.1NshRNA: n = 37 segments), but no differences in total spine length or head area. Bar graph (right)
shows no changes to the proportion of spine types between 4.1N-shRNA-expressing neurons
compared to GFP-expressing control neurons (GFP: n =5 cells, 4.1N-shRNA: n = 4 cells). (H)
AMPAR-mEPSC analysis reveals a significant reduction in the frequency, but not the amplitude,

38
of AMPAR-mEPSCs in 4.1N-shRNA-expressing DG granule neurons compared to control DG
granule neurons (control: n = 6 cells, 4.1N-shRNA: n = 6 cells). Bar graphs show the averaged
frequency and amplitude of AMPAR-mEPSC events ±SEM, with each point representing the
averaged value from one neuron. Leftmost panel shows sample traces of control AMPARmEPSC events (black/top), compared to 4.1N-shRNA AMPAR-mEPSC events (blue/bottom).
Scale bars: 1s, 20pA. Left of the amplitude bar graph displays an averaged representative trace
from a control (black) and transfected (blue) neuron. Scale bars: 5ms, 5pA. (I) Paired scatterplot
shows no differences in decay kinetics between averaged AMPAR-eEPSCs from 4.1N-shRNAexpressing and control DG granule neurons (n = 9 pairs). Inset shows peak-normalized sample
traces from control (black) and transfected (blue) neurons. Scale bar: 10ms. *p<0.05; n.s., Not
significant.

39
Figure 2.2 Protein 4.1N is not required for glutamatergic neurotransmission in CA1
pyramidal neurons.
(A) Representative immunolabeling showing minimal 4.1N expression in the CA1 region of a rat
hippocampal slice. Blue box shows enlarged CA1 region. SO: stratum oriens, SP: stratum
pyramidale, SR: stratum radiatum. Orange box shows enlarged DG region. GL: granule layer,
ML: molecular layer. (B) Schematic representation of electrophysiological recording setup for
CA1 pyramidal neurons. (C) Knockdown of 4.1N in CA1 pyramidal neurons does not
significantly affect AMPAR-eEPSC (n = 8 pairs) or (D) NMDAR-eEPSC amplitude (n = 8 pairs).
(C-D) Scatterplots show eEPSC amplitudes for pairs of untransfected and transfected cells (open
circles) with corresponding mean ±SEM (filled circles). Insets show representative current traces
from control (black) and transfected (blue) neurons with stimulation artifacts removed. Scale
bars: 20 ms, 20pA for AMPAR-eEPSCs. 50 ms, 50pA for NMDAR-eEPSCs. Bar graphs show
the average AMPAR-eEPSC and NMDAR-eEPSC amplitudes (±SEM) of CA1 pyramidal
neurons expressing the 4.1N-shRNA (blue) normalized to their respective control cell average

40
eEPSC amplitudes (black). (E) Paired-pulse facilitation ratios (mean ±SEM) for 4.1N-shRNAexpressing CA1 pyramidal neurons and paired control neurons show no detectable differences in
facilitation at a variety of interstimulus intervals (n = 6 pairs, ISIs: 20, 40, 70, and 100 ms). Peak
1- scaled current traces from control (black) and transfected (blue) neurons. Scale bars: 20 ms.
(F). Paired scatterplot shows no differences in decay kinetics between averaged AMPAR-eEPSCs
from 4.1N-shRNA-expressing and paired control neurons (n =7 pairs). Inset shows peaknormalized sample traces from control (black) and transfected (blue) neurons. Scale bar: 10ms.
n.s. Not significant.

41
Figure 2.3 Protein 4.1N’s C-terminal domain is not required for glutamatergic synapse
function in DG granule neurons.
(A) Schematic depicting the domain structure of full length 4.1N protein, followed by the domain
structure of 4.1N∆CTD. (B) Molecular replacement of endogenous 4.1N with shRNA-resistant
4.1N cDNA (4.1N Rescue) rescues both AMPAR-eEPSC (n = 9 pairs) and (E) NMDAR-eEPSC
amplitudes (n = 9 pairs) in DG granule neurons. Molecular replacement of endogenous 4.1N with
4.1N∆CTD also rescues both (C) AMPAR-eEPSC (n = 7 pairs) and (F) NMDAR-eEPSC
amplitudes (n = 7 pairs) in DG granule neurons. (B, C, E, F) Scatterplots show eEPSC amplitudes

42
for pairs of untransfected and transfected cells (open circles) with corresponding mean ±SEM
(filled circles). Insets show representative current traces from control (black) and transfected
(4.1N Rescue: gray, 4.1N∆CTD: red) neurons with stimulation artifacts removed. Scale bars: 20
ms, 20pA for both AMPAR-eEPSCs. 50ms, 20pA for 4.1N rescue NMDAR-eEPSCs. 20ms,
50pA for 4.1N∆CTD NMDAR-eEPSCs. (D, G) Bar graphs show the average AMPAR-eEPSC
and NMDAR-eEPSC amplitudes (±SEM) of DG granule neurons co-expressing: 4.1N-shRNA
and 4.1N-shRNA-resistant cDNA (4.1N Rescue: gray) or 4.1N-shRNA and 4.1N∆CTD (red)
normalized to their respective control cell average eEPSC amplitudes (black). (H) Paired
scatterplot shows no differences in the decay kinetics of averaged AMPAR-eEPSCs from
4.1N∆CTD-expressing compared to control DG granule neurons (n =6 pairs). Inset shows peaknormalized sample traces from control (black) and transfected (red) neurons. Scale bar: 10ms. (I)
Co-transfection of DG granule neurons with 4.1N-shRNA and 4.1N-shRNA-resistant cDNA
(4.1N Rescue) rescues the 4.1N-shRNA-mediated reduction in dendritic spine density. Leftmost
images display representative dendritic segments of GFP- (left), 4.1N-shRNA- (middle), and
4.1N Rescue (right) -expressing DG granule neurons. Scale bars: 10µm. Volcano plots show no
significant differences in dendritic spine density, spine length, or head area in DG granule
neurons co-expressing the 4.1N-shRNA with 4.1N-shRNA-resistant cDNA when compared to
GFP-expressing control neurons (GFP: n =31 segments, 4.1N Rescue: n = 48 segments). Bar
graph (right) shows no significant differences in proportion of spine types between neurons coexpressing 4.1N-shRNA and 4.1N-shRNA-resistant cDNA compared to GFP-expressing control
neurons (GFP: n =5 cells, 4.1N Rescue: n = 6 cells). *p<0.05; n.s., Not significant.

43
Figure 2.4 Protein 4.1N’s FERM domain is required for maintaining synaptic AMPA
receptor function in DG granule neurons.
(A) Schematic depicting the domain structure of 4.1N∆FERM. Molecular replacement of
endogenous 4.1N with 4.1N∆FERM rescues (D) NMDAR-eEPSC (n = 12 pairs) but (B) fails to
rescue AMPAR-eEPSC amplitudes (n = 13 pairs) in DG granule neurons. (B, D) Scatterplots
show eEPSC amplitudes for pairs of untransfected and transfected cells (open circles) with
corresponding mean ±SEM (filled circles). Insets show representative current traces from control
(black) and transfected (deep blue) neurons with stimulation artifacts removed. Scale bars: 20 ms,

44
20pA for AMPAR-eEPSCs. 50ms, 50pA for NMDAR-eEPSCs. (C, E) Bar graphs show the
average AMPAR- and NMDAR-eEPSC amplitudes (±SEM) of DG granule neurons coexpressing 4.1N-shRNA and 4.1N∆FERM (deep blue) normalized to their respective control cell
average eEPSC amplitudes (black). (F) Molecular replacement of endogenous 4.1N with
4.1N∆FERM rescues the 4.1N-shRNA-mediated reduction in dendritic spine density.
Representative dendritic segment of GFP- (left) and 4.1N-shRNA + 4.1N∆FERM (right) -
expressing DG granule neurons. Scale bars: 10µm. Volcano plots show no significant differences
in dendritic spine density, spine length, or head area in DG granule neurons co-expressing the
4.1N-shRNA with 4.1N∆FERM in comparison to GFP-expressing control neurons (GFP: n = 31
segments, 4.1N∆FERM: n = 31 segments). Bar graph (right) shows no significant differences in
proportion of spine types between neurons co-expressing 4.1N-shRNA and 4.1N∆FERM
compared to GFP-expressing control neurons (GFP: n = 5 cells, 4.1N∆FERM construct: n = 6
cells). (G) AMPAR-mEPSC analysis reveals a significant reduction in the frequency, but not the
amplitude, of AMPAR-mEPSCs in 4.1N∆FERM-expressing DG granule neurons compared to
control DG granule neurons (control: n = 6 cells, 4.1N∆FERM: n = 6 cells). Bar graphs show the
averaged frequency and amplitude of AMPAR-mEPSCs ±SEM, with each point representing the
averaged current of one neuron. Leftmost panel shows sample traces of control AMPAR-mEPSC
events (black/top), compared to 4.1N∆FERM AMPAR-mEPSC events (deep blue/bottom) Scale
bars: 500ms, 20pA. Left of the amplitude bar graph displays an averaged representative trace
from a control (black) and transfected (deep blue) neuron. Scale bars: 5ms, 2pA. (H) Coefficient
of variation analysis of AMPAR-eEPSCs from pairs of control and 4.1N∆FERM-expressing DG
granule neurons. Coefficient of Variation analysis reveals the reduction in AMPAR-eEPSC
amplitude caused by the loss of the FERM domain is due to reduction in quantal content (n=13

45
pairs). CV-2 values are plotted against corresponding ratios of mean amplitudes within each pair
(open circles) with mean ±SEM (filled circle). (I) Failure Analysis of AMPAR-eEPSCs reveals
that 4.1N∆FERM-expressing DG granule neurons exhibit significantly higher rates of failure
compared to control DG granule neurons (n= 13 pairs). (J) Paired scatterplot shows a significant
speeding in the decay kinetics of 4.1N∆FERM AMPAR-eEPSCs when compared to control DG
granule neurons (n =9 pairs). Inset shows peak-normalized sample traces from control (black) and
transfected (deep blue) neurons. Scale bar: 10ms. *p<0.05; n.s., Not significant.

46
Chapter 3: The Role of the 4.1 Family of Proteins in Hippocampal
Glutamatergic Synapse Regulation.
3.1 Abstract
The vertebrate 4.1 family of proteins is comprised of four evolutionarily conserved,
multifunctional proteins with variable expression patterns in the brain. We find family members
4.1R and 4.1B to be nonessential for regulating basal glutamatergic neurotransmission in both
perforant path- DG granule and Schaffer collateral –CA1 pyramidal synapses. Surprisingly, we
find loss of 4.1N’s spectrin-actin binding (SAB) domain dramatically augments AMPAR-eEPSC
amplitude in DG granule neurons. This AMPAR phenotype is the opposite of what is seen with
loss of 4.1N’s FERM domain at perforant path- DG granule synapses. This work highlights the
specificity between 4.1 proteins in hippocampal synapse regulation, while also providing
evidence that 4.1N is bidirectionally involved in maintaining stable levels of synaptic AMPAR
expression at DG granule synapses.
3.2 Introduction
Members of the 4.1 family of proteins are evolutionarily conserved, multifunctional,
membrane-cytoskeleton adapters (Baines et al., 2014). Vertebrates have four paralogues: 4.1R,
4.1N, 4.1B, and 4.1G. 4.1R, the prototypical family member, is a major mammalian erythrocyte
cytoskeletal protein (Correas et al., 1986). By forming a tripartite complex with spectrin and
actin, 4.1R maintains cytoskeletal stability in red blood cells. 4.1R is also critical for linking
membrane proteins to the actin cytoskeleton (Takakuwa, 2000).
4.1 proteins are defined by their domain structure (Fig. 3.1). All 4.1 proteins share two
domains: FERM and C-Terminal Domains (CTD). Both domains bind a variety of
transmembrane proteins. Vertebrate 4.1 proteins also share a FERM-adjacent (FA) domain, as

47
well as a spectrin-actin binding (SAB) domain. The FA domain functions to modulate the FERM
domain via regulatory phosphorylation sites (Baines et al., 2006). The SAB domain is responsible
for forming a ternary complex with spectrin and actin, thus stabilizing the erythrocyte
cytoskeleton in 4.1R (Gimm et al., 2002). Intriguingly, in vitro binding assays show 4.1B and
4.1G are less effective at interacting with actin to form a ternary complex. Further, 4.1N’s SAB
domain displays no binding affinity for spectrin or F-actin and is entirely unable to form this
complex (Gimm et al., 2002). Whether 4.1N’s SAB has mutated to bind other proteins or perform
another function remains to be seen.
Figure 3.1 Conserved domain structure of the protein 4.1 family.
Between the domains lie three regions of low complexity (U1, U2, U3). The N-Terminal
U1 region is sometimes referred to as the variably sliced headpiece (Conboy et al., 1988). Indeed,
alternative splicing of this region gives rise to the two main isoforms of 4.1R. While essential in
hematopoietic tissues, 4.1R’s role in other cell types is far less characterized. Outside of red blood
cells, in-situ hybridization shows 4.1R mRNA is selectively enriched in specific neuronal
populations within the cerebellum and DG (Walensky et al., 1998). 4.1R knockout mice show
deficits in movement, coordination, balance, and learning - in addition to the predicted
hematological abnormalities (Walensky et al., 1998). The 80kDA isoform of 4.1R, one of its two
most abundant isoforms, is enriched 11-fold in postsynaptic densities (Scott et al., 2001). In fact,
all 4.1 proteins bind a diverse array of synaptic proteins and display significant enrichment in

48
postsynaptic densities - suggesting they may be involved in regulating synaptic neurotransmission
(Scott et al., 2001).
Figure 3.2 Hippocampal mRNA expression varies between members of the 4.1 family of
proteins (Figure adapted from Parra et al., 2000).
Despite retaining significant structural homology, 4.1 proteins display distinct expression
patterns. In-situ hybridization shows 4.1B, 4.1N, and 4.1R are significantly enriched in disparate
hippocampal subregions (Fig. 3.2; adapted from Parra et al., 2000). While 4.1R mRNA is
concentrated in DG granule neurons, 4.1B mRNA is localized to pyramidal neurons of CA3 and
CA1 (Parra et al., 2000). The diverse expression levels in the hippocampus between these
proteins suggest differences in binding partners and regulation of activity. 4.1G mRNA, on the
other hand, is highly concentrated in the retina – with far lower levels seen in the cerebrum and
other brain regions (Sanuki et al., 2015). As such, we sought to assess a potential synaptic role for
two 4.1 proteins, 4.1B and 4.1R, with hippocampal-specific mRNA expression (Parra et al.,
2000). We find knockdown of 4.1R does not disrupt basal glutamatergic neurotransmission in
either Schaffer collateral- CA1 pyramidal or perforant path – DG granule synapses. Similarly, we
find knockdown of 4.1B does not disrupt basal glutamatergic neurotransmission in these two
neuronal populations. Next, we examined the functional effects of removing 4.1N’s SAB domain.

49
Removal of 4.1N’s SAB domain produces a dramatic increase in synaptic AMPAR currents in
DG granule neurons, the opposite of what is seen with loss of 4.1N’s FERM domain. This work
contributes to the growing body of literature detailing the molecular specificity found in synaptic
regulation. Further, we provide evidence that protein 4.1N bidirectionally maintains stable
synaptic AMPAR expression in DG granule neurons.
3.3 Materials & Methods
Experimental Constructs: A previously validated 4.1R-shRNA target sequence against 4.1R was
used for all 4.1R knockdown experiments (5’- CCAGACATGTCAGTGACCA-3’; Huang et al.,
2016). A previously validated 4.1B-shRNA target sequence was used for all 4.1B knockdown
experiments (5’-GCAGGGGCACAGGCTGCACCT -3’). Both the 4.1R-shRNA and the 4.1BshRNA were independently subcloned behind the H1 promoter region of a GFP-expressing
pFHUGW expression vector. The shRNA-resistant 4.1NΔSAB mutant was generated by deleting
the residue corresponding to the SAB domain (Thr477- Ala543) and was obtained from Genscript
(Cat. #SC1626). The shRNA-resistant 4.1NΔSAB mutant was cloned into a pCAGGS-IRESmCherry expression vector. All plasmids were confirmed by DNA sequencing. A pFUGW vector
expressing only GFP was co-expressed with pCAGGS-IRES-mCherry constructs to enhance
identification of transfected neurons and was also used as a control vector in spine imaging
experiments.
Electrophysiology: All experiments were performed in accordance with NIH Guidelines
for the Care and Use of Laboratory Animals, and all procedures were approved by the
Institutional Animal Care and Use Committee of the University of Southern California. 400 μm
rat organotypic entorhino-hippocampal slice cultures were prepared from both male and female
post-natal day 6-8 (P6-8) Sprague Dawley rats as previously described (Stoppini et al., 1991;

50
Prang et al., 2001; Bonnici and Kapfhammer, 2009; Sadybekov et al., 2017; Tian et al., 2018).
Tissue was isolated and a MX-TS tissue slicer (Siskiyou) was used to make 400 μm transverse
sections. Tissue slices were placed on squares of Biopore Membrane Filter Roll (Millipore) and
placed on Millicell Cell Culture inserts (Millipore) in 35 mm dishes. Slices were fed 1mL of
culture media containing MEM + HEPES (Gibco Cat#12360-038), horse serum (25%), HBSS
(25%) and L-glutamine (1 mM). Media was exchanged every other day. Slices with large portions
of entorhinal cortex were visually identified after slicing. These slices were selected and plated
for use in our experiments, and presence of entorhinal cortex was again confirmed when selecting
slices appropriate for data acquisition. Whole-cell recordings were performed on day in vitro 7-8
(DIV7-8). During recordings, slices were maintained in room-temperature artificial cerebrospinal
fluid (aCSF) containing 119 mM NaCl, 2.5 mM KCl, 1 mM NaH2PO4, 26.2 mM NaHCO3 11 mM
glucose, 4 mM CaCl2, and 4 mM MgSO4. 5 μM 2-chloroadenosine and 0.1 mM picrotoxin were
also added to the aCSF to dampen epileptiform activity and block GABAA receptor activity,
respectively. Osmolarity was adjusted to 310-315 mOsm. aCSF was saturated with 95% O2/5%
CO2 throughout the recording. Borosilicate recording electrodes were filled with an internal
whole-cell recording solution containing 135 mM CsMeSO4, 8 mM NaCl, 10 mM HEPES, 0.3
mM EGTA, 5 mM QX-314, 4 mM Mg-ATP, and 0.3 mM Na-GTP. Osmolarity was adjusted to
290–298 mOsm, and pH-buffered at 7.3–7.4.
DG granule neurons and CA1 pyramidal neurons were identified using differential
interference phase contrast microscopy, while GFP-expressing transfected neurons were
identified using epifluorescence microscopy. Dual whole-cell recordings of either neuronal
subtype were made through simultaneous recordings from a transfected neuron and a
neighboring, untransfected control neuron. Synaptic responses were evoked by stimulating with a

51
monopolar glass electrode filled with aCSF in the stratum radiatum for CA1 recordings and the
perforant pathway for DG granule neuron recordings. Membrane holding current, pipette series
resistance, and input resistance were monitored throughout recording sessions. Data were
acquired using a Multiclamp 700B amplifier (Molecular Devices), filtered at 2 kHz, and digitized
at 10 kHz. AMPAR-evoked EPSCS (-eEPSCs) were measured at -70mV. NMDAR-eEPSCs were
measured at +40 mV and were temporally isolated by measuring amplitudes 150 ms following the
stimulus, at which point the AMPAR-eEPSC has completely decayed. Data analysis was
performed using Igor Pro (Wavemetrics). In the scatter plots for simultaneous dual whole-cell
recordings, each open circle represents one paired recording, and the closed circle represents the
average of all paired recordings. No more than one paired recording was performed on any given
entorhino-hippocampal slice.
Biolistic Transfection: Sparse biolistic transfections were performed on DIV1 as
previously described (Stoppini et al., 1991; Schnell et al., 2002; Lu et al., 2009). 50μg of mixed
plasmid DNA was coated on 1μm-diameter gold particles in 0.5 mM spermidine, precipitated
with 0.1 mM CaCl2, and washed four times in pure ethanol. The DNA-coated gold particles were
then coated onto PVC tubing, dried briefly using ultra-pure N2 gas, and stored at 4 °C in
desiccant. Before use, the gold particles were brought up to room temperature and delivered to
slice cultures via a Helios Gene Gun (BioRad). Construct expression was confirmed by GFP or
mCherry epifluorescence.
Spine Density Analysis: Cultured entorhino-hippocampal slices were transfected on DIV1
with pFUGW-GFP construct, pFHUGW-GFP-shRNA construct, or pFHUGW-GFP-shRNA +
pCAGGS-mCherry-cDNA constructs. Slices were fixed in 4% PFA, 4% sucrose in PBS, and
washed three times with PBS, then cleared with an abbreviated SeeDB-based protocol (Ke et al.,

52
2013) and mounted on microscope slides. Images were acquired at DIV7 using super-resolution
microscopy (Carl Zeiss). High-resolution confocal z-stacks of spine-containing DG granule
neuron secondary apical dendrites were acquired on a Zeiss 880 using an EC Plan-Neofluar
40x/1.3 oil-immersion DIC M27 objective. ~60 μm sections of secondary apical dendrites were
manually selected for analysis. Z-stacks were collected at maximum X–Y pixel dimensions (512
x 512 pixels) at 8 bits with a 488 nm laser excitation wavelength. An experimenter, blinded to the
experimental condition, performed spine density analysis on sections using the Dendritic Spine
Counter plug-in on ImageJ to count spines extending laterally from the dendrite. The ImageJ
plug-in Dendritic Spine Counter was also used to obtain values for the following spine
morphology metrics: spine neck length, neck width, and head width (used for spine type
classification). Dendritic processes are commonly classified into the following categories: stubby,
mushroom, and thin. Based on previous literature, we categorized dendrites into three
aforementioned categories using the following criteria. Stubby spines are those which lack a
visible neck region but have a bulbous head. Mushroom spines, often viewed as mature spines,
are those with a short length (<2um), a clearly defined neck region, and a head which is >50% as
wide as the neck. Thin spines are those with a long length (>2um) as well as a head (Mattison et
al., 2014; Lin et at., 2004).
Experimental Design and Statistical Analyses: All electrophysiological data are expressed
as mean ± standard error measurement (SEM). Imaging analysis was performed blind to
experimental condition. Statistical significance was determined using Wilcoxon signed-rank test
for paired dual whole-cell patch clamp data and Wilcoxon rank-sum test for imaging data. Data
were analyzed using IGOR Pro (Wavemetrics RRID:SCR_000325) or KaleidaGraph (Synergy
Software RRID:SCR_014980) and graphed using Microsoft Excel (RRID:SCR_016137) or

53
GraphPad Prism (RRID:SCR_002798). All p-values <0.05 were considered significant and
denoted with a single asterisk. All error bars represent standard error measurement. For all
experiments, at least 4 male and female rat pups were used. Sample sizes in the present study are
similar to those reported in the literature (Herring and Nicoll, 2016; Incontro et al., 2018).
3.4 Results
4.1R knockdown does not perturb basal glutamatergic synapse function in CA1 pyramidal
neurons or DG granule neurons.
Utilizing a previously published RNAi sequence for 4.1R, we generated a 4.1R-shRNA contruct
using an in-house vector (Huang et al., 2016). Following this, we employed a biolistic
transfection method to express the 4.1R-shRNA in neurons of rat organotypic entorhinohippocampal slice cultures (Elias et al., 2008; Paskus et al., 2019; Tian et al., 2021). These slices
provide a method to examine the impact a genetic modification has on hippocampal neurons
within their native circuitry (Stoppini et al., 1991; Schnell et al., 2002). 6 days after transfection,
we record AMPA- and NMDA-receptor evoked excitatory postsynaptic currents (AMPAR- and
NMDAR-eEPSCs) from transfected and neighboring, untransfected control neurons
simultaneously during Schaffer collateral (Fig 3.3B) or perforant pathway stimulation (Fig. 3.3C).
This approach allows for a pairwise, internally controlled comparison of the consequences of our
acute genetic manipulation. We find that knockdown of protein 4.1R in CA1 pyramidal neurons
produces no significant change to either AMPAR-eEPSC amplitude (n = 8 pairs, p = 0.46,
Wilcoxon signed-rank test; Fig. 3.3D, F) or NMDAR-eEPSC amplitude (n = 7 pairs, p = 0.11,
Wilcoxon signed-rank test; Fig. 3.3G, I). Similarly, we find knockdown of protein 4.1R in DG
granule neurons does not alter either AMPAR-eEPSC amplitude (n = 8 pairs, p = 1, Wilcoxon

54
signed-rank test; Fig. 3.3E, F) or NMDAR-eEPSC amplitude (n = 8 pairs, p = 0.64, Wilcoxon
signed-rank test; Fig. 3.3H, I). These data strongly suggest that 4.1R has no role in regulating
basal synaptic transmission in either CA1 pyramidal or DG granule neurons.
4.1B knockdown does not perturb basal glutamatergic synapse function in CA1 pyramidal
neurons or DG granule neurons.
Using the method detailed above, we generated a 4.1B-shRNA construct and knocked down
protein 4.1B via biolistic transfection into organotypic entorhino-hippocampal slice cultures. We
find that knockdown of protein 4.1B in CA1 pyramidal neurons produces no significant change to
either AMPAR-eEPSC amplitude (n = 8 pairs, p = 0.84, Wilcoxon signed-rank test; Fig. 3.4D, F)
or NMDAR-eEPSC amplitude (n = 8 pairs, p = 1, Wilcoxon signed-rank test; Fig. 3.4G, I).
Similarly, we find knockdown of protein 4.1B in DG granule neurons results in no alterations to
AMPAR-eEPSC amplitude (n = 5 pairs, p = 0.81, Wilcoxon signed-rank test; Fig. 3.4E, F) or
NMDAR-eEPSC amplitude (n = 5 pairs, p = 1, Wilcoxon signed-rank test; Fig. 3.4H, I). Taken
together, we conclude that 4.1B has no role in regulating basal glutamatergic neurotransmission
in either CA1 pyramidal or DG granule neurons of the hippocampus.
Loss of 4.1N’s SAB domain augments synaptic AMPAR currents in DG granule neurons.
Canonically, the SAB domain of 4.1 proteins is responsible for forming a ternary complex with
spectrin and actin (Gimm et al., 2002). Intriguingly, in vitro binding assays show 4.1N’s SAB
domain has no binding affinity for spectrin or F-actin and is entirely unable to form this complex
(Gimm et al., 2002). Whether 4.1N’s SAB domain has mutated to bind other proteins or perform
novel functions remains to be seen. To determine the role of protein 4.1N’s SAB domain in

55
regulating DG granule synapses, we molecularly replaced endogenous 4.1N with a mutant variant
lacking its SAB domain (4.1NΔSAB; Fig. 3.5A). We find that molecular replacement of
endogenous 4.1N with 4.1NΔSAB rescues the reduction in NMDAR-eEPSC amplitude caused by
4.1N knockdown (n = 9 pairs, p = 0.36, Wilcoxon signed-rank test; Fig. 3.5D). Surprisingly,
4.1NΔSAB fails to rescue the reduction in AMPAR-eEPSC amplitude caused by 4.1N
knockdown (n = 9 pairs, p = 0.0039, Wilcoxon signed-rank test; Fig. 3.5C). Instead, we observe
a significant increase in synaptic AMPAR currents in 4.1NΔSAB-expressing DG granule
neurons. Due to this unexpected phenotype, we assessed whether removal of 4.1N’s SAB domain
is accompanied with a change in the number or morphology of dendritic spines at DG granule
synapses. We find that co-expression of 4.1N-shRNA with 4.1NΔSAB cDNA results in a
significant increase in dendritic spine density (GFP: n =17 segments, 4.1N∆SAB: n = 21
segments, p < 0.0001, Wilcoxon rank-sum test; Fig. 3.5E). Despite the observed change in spine
density, we find the proportion of spine types in DG granule neurons co-expressing 4.1N-shRNA
and 4.1N∆SAB compared to GFP-expressing control neurons to remain unaltered (GFP: n = 5
cells, 4.1N∆SAB: n = 4 cells, p = 0.56 for stubby spines, p = 1 for thin spines, p = 0.41 for
mushroom spines, Wilcoxon rank-sum test; Fig. 3.5E).
3.5 Discussion
The vertebrate 4.1 family of proteins is comprised of four highly conserved,
multifunctional, membrane-cytoskeleton adapters. Despite their similarities, these proteins show
variable expression patterns in the brain. Previous immunohistochemical data from our lab shows
protein 4.1N is expressed in a site-specific manner within the hippocampus. Protein 4.1N shows
robust dendritic immunoreactivity in DG granule neurons. In contrast, 4.1N has considerably
lower levels of expression in the dendrites of CA3 and CA1 pyramidal neurons. This pattern of

56
expression is distinct from other members of the 4.1 family. Within the brain, 4.1G is highly
expressed in the retina (Sanuki et al., 2015). 4.1B mRNA, on the other hand, shows high levels of
expression in cerebellar Purkinje cells and hippocampal pyramidal neurons (Parra et al., 2000).
4.1R also displays discrete patterns of localization, with hippocampal expression restricted to
granule cells of the dentate gyrus (Walensky et al., 1998; Parra et al., 2000). As such, we
investigated whether 4.1 proteins with site-specific hippocampal expression had any role in
synaptic regulation within CA1 pyramidal or DG granule neurons.
We find knockdown of protein 4.1R in either CA1 pyramidal or DG granule neurons does
not perturb basal glutamatergic neurotransmission. Protein 4.1R, the canonical 4.1 family
member, is essential for stabilizing the erythrocyte cytoskeleton (Takakuwa, 2000). By forming a
tripartite complex with spectrin and actin, 4.1R maintains membrane stability. As such, 4.1R
plays a role in the immunological synapse. Within the brain, protein 4.1R expression is restricted
to granule neurons of the cerebellum and DG (Walensky et al., 1998). Mice lacking 4.1R show
predicted hematological abnormalities, as well as deficits in moving, coordination, and balance
(Walensky et al., 1998; Shi et al., 1999). These latter functions are attributable to the cerebellum,
in which granule cells are the only excitatory cell type and comprise over 99% of total cerebellar
neurons (Herculano-Houzel et al., 2006; Consalez et al., 2020). It stands that 4.1R, in a manner
similar to 4.1N, may possess a site-specific role in synaptic transmission within granule neurons
of the cerebellum. Future work may wish to examine the functional effects of an acute
knockdown within these synapses.
We also find that reducing protein 4.1B expression has no effect on altering glutamatergic
neurotransmission in either CA1 pyramidal and DG granule neurons. Previous research has
uncovered that SynCAM1, a synaptic adhesion molecule, recruits NMDARs via protein 4.1B

57
(Biederer et al., 2002; Hoy et al., 2009). In conjunction with SynCAM1, 4.1B increases the
frequency of NMDAR-mediated mEPSCs in HEK293/neuronal co-culture assays (Hoy et al.,
2009). This interaction was examined in vitro, utilizing dissociated assays in which determining
neuronal identity is difficult. As such, it remains possible that 4.1B and SynCAM1 have a similar
role in vivo in a different brain region. Recent work has shown an in vivo role for 4.1B in proper
synapse formation in the developing zebrafish spinal cord (Fierro et al., 2018). Within the
zebrafish, 4.1B has two isoforms with distinct patterns of expression. Knockdown of one isoform
reduces the number of glutamatergic synapses in motor neurons (Fierro et al., 2018). If this role is
conserved in humans, it stands that 4.1B may be critical for synaptic function in motor neurons.
Going forward, it will be important to carry out thorough investigations of 4.1B’s role in synaptic
regulation within cortical motor neurons.
Previously, we discovered that knocking down 4.1N in DG granule neurons results in a
reduction in AMPAR-and NMDAR-eEPSC amplitudes that is caused by a loss of dendritic
spines. We also show that 4.1N’s FERM domain is essential for supporting synaptic AMPAR
function in these neurons. However, the domain(s) required for 4.1N’s role in supporting
glutamatergic synapse structure remains unknown. Initially, we hypothesized that 4.1N’s SAB
domain promotes the recruitment of actin regulatory proteins that are required for the
maintenances of dendritic spines. Canonically, 4.1R’s SAB domain binds spectrin and actin to
form a tripartite complex essential for stabilizing the erythrocyte cytoskeleton (Takakuwa, 2000).
Biochemical studies show 4.1N’s SAB domain, on the other hand, is no longer able to form this
complex. Further, 4.1N’s SAB domain displays no binding affinity for spectrin or actin at all
(Gimm et al., 2002). Additional research on 4.1N’s SAB domain and novel binding partners has
been scarce. As such, it remains plausible that 4.1N’s SAB domain has mutated to bind other

58
synaptic proteins. We examined whether 4.1N’s SAB domain has a role in regulating perforant
path-DG granule synapses. Unexpectedly, we find molecular replacement of endogenous 4.1N
with 4.1N∆SAB dramatically augments AMPAR-eEPSC amplitude. This data suggests 4.1N is
bidirectionally involved in maintaining stable levels of synaptic AMPAR expression at DG
granule synapses. Whereas disruption of 4.1N’s FERM domain increases the number of synapses
lacking functional AMPARs, disruption of 4.1N’s SAB domain produces an increase in synaptic
AMPAR function. In this way, the FERM domain helps maintain stable synaptic AMPAR
expression – while the SAB domain acts as a molecular break on glutamatergic synapse strength.
Previous work from our laboratory uncovered a site-specific role for βSAP97 in
augmenting synaptic AMPAR-eEPSC amplitude (Kay et al., 2022). Reducing βSAP97 in DG
granule neurons significantly increases synaptic AMPAR currents, similar to what we find with
loss of 4.1N’s SAB domain. βSAP97 binds 4.1N via its I3 insert, and the two are
developmentally regulated in a similar fashion to GluA1 (Douyard 2007; Kay et al 2022). The
4.1N domain which βSAP97 binds to is not yet experimentally known. As such, it remains
possible that 4.1N’s SAB domain has gained the ability to bind βSAP97. Going forward, it will
be of great interest to examine the 4.1N domain responsible for this protein binding. Further, it
will be critical to experimentally assess whether 4.1N and βSAP97 are involved in the same
molecular pathway maintaining stable synaptic AMPAR expression at DG granule synapses.
In this work, we examined a potential synaptic role for 4.1B and 4.1R, two 4.1 proteins
with hippocampal-specific mRNA expression (Parra et al., 2000). We find knockdown of 4.1R
does not disrupt basal glutamatergic neurotransmission in either Schaffer collateral- CA1
pyramidal or perforant path – DG granule synapses. Similarly, we find knockdown of 4.1B does
not perturb basal glutamatergic neurotransmission in these neuronal populations. We also

59
examined the role of 4.1N’s SAB domain in synaptic regulation of DG granule neurons.
Surprisingly, we find removal of 4.1N’s SAB domain starkly increases synaptic AMPAR
currents. This phenotype is the opposite of what is seen with loss of 4.1N’s FERM domain at DG
granule synapses. Taken together, this work adds to the growing body of evidence detailing the
molecular specificity of synaptic regulation in different neuronal populations. Further, we present
evidence that 4.1N, a protein with a site-specific role in regulating glutamatergic
neurotransmission, provides both an upper and lower limit on synaptic AMPAR function in DG
granule neurons.

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Figure 3.3 Knockdown of protein 4.1R does not disrupt basal glutamatergic
neurotransmission in either CA1 pyramidal or DG granule synapses.

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(A) Conserved domain structure of vertebrate 4.1 proteins. (B) Schematic representation of
electrophysiological recording setup for CA1 pyramidal neurons. (C) Schematic representation of
electrophysiological recording setup for DG granule neurons. (D) Knockdown of 4.1R in CA1
pyramidal neurons does not significantly affect AMPAR-eEPSC (n = 8 pairs) or (G) NMDAReEPSC amplitudes (n = 7 pairs). (E) Knockdown of 4.1R in DG granule neurons does not
significantly alter AMPAR-eEPSC (n = 8 pairs) or (H) NMDAR-eEPSC amplitudes (n = 8 pairs).
(D, E, G, H) Scatterplots show eEPSC amplitudes for pairs of untransfected and transfected cells
(open circles) with corresponding mean ±SEM (filled circles). Insets show representative current
traces from control (black) and transfected (light blue, dark blue) neurons with stimulation
artifacts removed. Scale bars: 10 ms, 10pA for CA1 AMPAR-eEPSCs. 20ms, 10pA for DG
AMPAR-eEPSCs. 20ms, 10pA for both CA1 and DG NMDAR-eEPSCs. (F, I) Bar graphs show
the average AMPAR-eEPSC and NMDAR-eEPSC amplitudes (±SEM) of neurons expressing the
4.1R-shRNA (light blue, dark blue) normalized to their respective control cell average eEPSC
amplitudes (black).

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Figure 3.4 Knockdown of protein 4.1B does not disrupt basal glutamatergic
neurotransmission in either CA1 pyramidal or DG granule synapses.

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(A) Conserved domain structure of vertebrate 4.1 proteins. (B) Schematic representation of
electrophysiological recording setup for CA1 pyramidal neurons. (C) Schematic representation of
electrophysiological recording setup for DG granule neurons. (D) Knockdown of 4.1B in CA1
pyramidal neurons does not significantly alter AMPAR-eEPSC (n = 8 pairs) or (G) NMDAReEPSC amplitudes (n = 8 pairs). (E) Knockdown of 4.1B in DG granule neurons does not
significantly affect AMPAR-eEPSC (n = 5 pairs) or (H) NMDAR-eEPSC amplitudes (n = 5
pairs). (D, E, G, H) Scatterplots show eEPSC amplitudes for pairs of untransfected and
transfected cells (open circles) with corresponding mean ±SEM (filled circles). Insets show
representative current traces from control (black) and transfected (pink, purple) neurons with
stimulation artifacts removed. Scale bars: 10ms, 20pA for CA1 AMPAR-eEPSCs. 10ms, 10pA
for CA1 NMDAR-eEPSCs. 10ms, 10pA for DG AMPAR-eEPSCs. 20ms, 20pA for DG
NMDAR-eEPSCs. (F, I) Bar graphs show the average AMPAR-eEPSC and NMDAR-eEPSC
amplitudes (±SEM) of neurons expressing the 4.1B-shRNA (pink, purple) normalized to their
respective control cell average eEPSC amplitudes (black).

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Figure 3.5 Loss of protein 4.1N’s SAB domain augments synaptic AMPAR currents in DG
granule neurons.

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(A) Schematic depicting the domain structure of 4.1N∆SAB. (B) Schematic representation of
electrophysiological recording setup for DG granule neurons. Molecular replacement of
endogenous 4.1N with 4.1N∆SAB rescues (D) NMDAR-eEPSC amplitudes (n = 9 pairs) and (C)
produces a dramatic increase in AMPAR-eEPSC amplitudes (n = 9 pairs) in DG granule neurons.
(C, D) Scatterplots show eEPSC amplitudes for pairs of untransfected and transfected cells (open
circles) with corresponding mean ±SEM (filled circles). Insets show representative current traces
from control (black) and transfected (violet) neurons with stimulation artifacts removed. Scale
bars: 20 ms, 20pA for AMPAR-eEPSCs. 20ms, 40pA for NMDAR-eEPSCs. (C, D) Bar graphs
show the average AMPAR- and NMDAR-eEPSC amplitudes (±SEM) of DG granule neurons coexpressing 4.1N-shRNA and 4.1N∆SAB (violet) normalized to their respective control cell
average eEPSC amplitudes (black). (E) Molecular replacement of endogenous 4.1N with
4.1N∆SAB significantly increases dendritic spine density. Representative dendritic segment of
GFP- (left) and 4.1N-shRNA + 4.1N∆SAB (right) -expressing DG granule neurons. Scale bars:
10µm. Volcano plots show a significant increase in the dendritic spine density of DG granule
neurons co-expressing the 4.1N-shRNA with 4.1N∆SAB in comparison to GFP-expressing
control neurons (GFP: n = 17 segments, 4.1N∆SAB: n = 21 segments). Bar graph (right) shows
no significant differences in proportion of spine types between neurons co-expressing 4.1NshRNA and 4.1N∆SAB compared to GFP-expressing control neurons (GFP: n = 5 cells,
4.1N∆SAB construct: n = 4 cells).

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CHAPTER 4: CONCLUSION
Neurons, specialized cells within the central nervous system, transmit information at
specialized sites called synapses. The formation and maintenance of a synapse is controlled by a
wide array of selectively expressed neuropeptides and molecules (Lüscher et al., 2000; Huganir
and Nicoll, 2013; Südhof, 2018). Historically, methodological limitations have confined research
into synaptic function to one cell type. Recent technological advances in techniques such as
whole brain synaptome cartography and 3D electron microscopy have allowed researchers to
delve into the vast molecular differences between brain region-specific synapse subtypes (Roy et
al., 2018; Cizeron et al., 2020; Zhu et al., 2021). Single-cell transcriptomics and proteomic massspectrometry have provided further evidence for the immense heterogeneity between neuronal
cell types. Differences in connectivity, neuropeptide expression, and firing patterns all contribute
to a complex neural network able to modulate itself rapidly and accordingly in response to
constant sensory input (Ayhan et al., 2021; Zeisel et al., 2018). Despite these advances, more
research needs to be done to truly grasp the extent of these differences.
Investigations into synaptic function often examine Schaffer collateral-CA1 synapses of
the hippocampus. Findings from these studies are then frequently extrapolated to be true for all
synapses in the brain. In contrast to heavily studied Schaffer collateral- CA1 synapses, much less
is known about perforant path – DG granule synapses. Glutamatergic synapses onto DG granule
neurons are the gateway for information flow into the hippocampus. Regulation of the number,
size, and strength of these synapses contributes to the DG’s ability to perform critical functions,
including pattern separation (Leutgeb et al., 2007; McHugh et al., 2007). Deficits in pattern
separation, as well as overall dysfunction within the dentate gyrus, are heavily associated with the

67
development of complex brain disorders such as schizophrenia (Das et al., 2014; Yun et al.,
2016).
Growing evidence suggests perforant path-DG granule synapses are regulated by a unique
set of synaptic proteins. One such molecular regulator is the synaptic scaffolding protein SAP97.
Work from our lab has recently uncovered a site-specific role for βSAP97 in augmenting synaptic
AMPAR-eEPSC amplitude in DG granule neurons (Kay et al., 2022). Immunohistochemistry
shows βSAP97 expression is concentrated in the dendrites of DG granule neurons, with
significantly lower levels of expression in CA1 and CA3 pyramidal neurons. We find βSAP97
sequesters perisynaptic GluA1-containing AMPARs away from perforant pathway- DG granule
synapses, a function it does not perform in Schaffer collateral - CA1 synapses.
Another protein of interest in synaptic regulation is 4.1N. Despite binding many
glutamatergic synapse proteins, 4.1N’s role at synapses has long eluded researchers. Chapter 2
describes a novel, postsynaptic, site-specific role for protein 4.1N in regulating perforant
pathway- DG granule synapses of the hippocampus. We observe robust dendritic protein 4.1N
immunoreactivity in DG granule neurons, with much lower protein expression in the dendrites of
CA3 and CA1 pyramidal neurons. Inhibition of protein 4.1N in DG granule neurons markedly
disrupts both AMPA- and NMDA-receptor mediated neurotransmission. We find this role is
partially dependent on 4.1N’s FERM domain, as opposed to its well-studied CTD. Removing
4.1N’s FERM domain increases the number of silent synapses, synapses which lack functional
AMPARs. Further, loss of 4.1N’s FERM domain phenocopies the synaptic alterations observed
when GluA1-containing AMPARs are absent from neurons. Our results strongly suggest that
4.1N’s FERM domain plays a critical role in GluA1-containing AMPAR trafficking to DG
granule synapses.

68
Chapter 3 introduces a second role for protein 4.1N in synaptic regulation. Unexpectedly,
loss of 4.1N’s SAB domain dramatically augments synaptic AMPAR function in DG granule
neurons. This data suggests 4.1N is bidirectionally involved in maintaining stable levels of
synaptic AMPAR expression at DG granule synapses. Whereas disruption of 4.1N’s FERM
domain decreases the number of synapses containing functional AMPARs, loss of 4.1N’s SAB
domain increases synaptic AMPAR function. In this way, the FERM domain helps maintain
stable synaptic AMPAR expression – while the SAB domain acts as a molecular break on
glutamatergic synapse strength.
This work demonstrates a major role for protein 4.1N in maintaining basal glutamatergic
synapse structure and function. Further, this role is cell-type specific within the hippocampus.
4.1N joins a growing list of synaptic proteins with region-specific roles in regulating perforant
path- DG granule synapses (Roy et al., 2018; Rao et al., 2019; Grant et al., 2020; Kay et al.,
2022). We believe protein 4.1N acts as a master regulator of synaptic development in DG granule
neurons, contributing to the hypothesis that DG granule synapses are controlled by a unique set
of peptides and molecules.
Chapter 3 also investigates potential roles for 4.1R and 4.1B, other 4.1 family members
with localized hippocampal expression, in synaptic regulation. We find that neither 4.1R nor
4.1B have significant roles in regulating glutamatergic neurotransmission at Schaffer collateralCA1 or perforant path-DG granule synapses. Nevertheless, it remains possible that these proteins
aid synaptic function in other brain regions. The work in this dissertation has only examined
differences between two types of synapses – both found within the hippocampus. Previous
research confirms there are significant differences between the proteins critical for synaptic
function in varying brain areas, even when neuronal types appear outwardly similar (Zeisel et al.,

69
2018; van Oostrum et al., 2023). Molecular specificity in synaptic regulation increases
informational complexity, aiding complex brain functions such as learning and memory.
Our understanding of the hippocampus’ contributions to learning, memory, and other
behaviors has grown vastly in the past 60 years. Nevertheless, many of its complexities remain a
mystery. Different subregions and neuronal populations contribute to synaptic function in
varying ways, made evident through lesion and molecular perturbation studies. Going forward, it
will be of great importance to understand how the unique proteins present at perforant pathway
synapses support information processing in the dentate gyrus. This knowledge will inform how
deficits in the functionality of these proteins contributes to the development of complex brain
disorders. My hope is that the work in this dissertation helps enhance our understanding of the
complex molecular mechanisms underlying glutamatergic synapse function.

70
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APPENDIX: Investigating a postsynaptic role for neurexins.
5.1 Introduction
Neurexins belong to a well-studied family of cell adhesion proteins. Initially, neurexins
were discovered as a receptor for α-Latrotoxin -a neurotoxin which produces immense
neurotransmitter release from presynaptic nerve terminals (Ushkaryov et al., 1992; Ushkaryov et
al., 2008). Three genes (NRXN1, NRXN2, NRXN3) comprise this family of cell adhesion
molecules in mammals, each of which undergoes extensive alternative splicing to create
thousands of transcript variants (Ushkaryov et al., 1992, Ullrich et al., 1995). Each neurexin gene
contains independent alternative promoters. These promoters drive expression of a long α
isoform, as well as a shorter β isoform (Chih et al., 2006; Missler and Südhof, 1998). Recent
work has discovered an even shorter γ isoform in mouse NRXN1, with an ortholog discovered in
C.elegans (Yan et al., 2015; Kurshan et al., 2018).
Figure 5.1 Domain structure of α- and β-neurexins.
Figure 5.1 displays the conserved domain structure of α- and β-neurexin isoforms. αneurexins contain six extracellular laminin/neurexin/sex hormone–binding globulin (LNS)
domains. Between the LNS domains are three epidermal growth factor (EGF)-like repeats.
Following the sixth LNS domain, there is a highly O-linked glycosylated stalk. After the stalk
comes the transmembrane region, followed by a cytoplasmic tail containing a 4.1 binding domain
and PDZ binding motif (Reissner et al., 2013; Trotter et al., 2019). The shorter β isoform lacks all

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EGF-like repeats and contains only the sixth extracellular LNS domain, the glycosylated stalk,
transmembrane region, and intracellular tail domain (Ushkaryov et al., 1994; Yan et al., 2015).
The NRXN1γ isoform is shorter still, retaining only the transmembrane region and cytoplasmic
tail (Yan et al., 2015).
Neurexins are presynaptic receptors for many extracellular and postsynaptic binding
partners, including neuroligins, cerebellins, dystroglycans, leucine-rich repeat transmembrane
proteins (LRRTMs), and more (Ichtchenko et al., 1995; Uemura et al., 2010; Sugita et al., 2001;
de Wit et al., 2009). Extensive alternative splicing modulates the binding affinity of individual
isoforms for many postsynaptic partners, including cerebellins, neuroligins, and latrophilins
(Boucard et al., 2005; Craig and Kang, 2007; Boucard et al., 2014). Alternative splicing of site
four also augments the mobility of both α- and β-isoforms, altering their functional properties
within synapses (Neupert et al., 2015). These differences contribute to the vast diversity seen
between synapse subtypes (Südhof 2017; Gomez et al., 2021).
While copious literature suggests neurexin expression is largely presynaptic, there is
evidence for postsynaptic localization of neurexins (Peng et al., 2004; Kattenstroth et al., 2004;
Taniguchi et al., 2007). Neurexins are enriched in postsynaptic densities, and cis-expressed βneurexins inactivate neuroligin-1 and promote the destabilization of synapses (Peng et al., 2004;
Taniguchi et al., 2007). Another study found that postsynaptic NMDAR function requires αneurexins (Kattenstroth et al., 2004). Despite the tremendous progress made in understanding the
function of these cell adhesion molecules, more work needs to be done to truly grasp the vast
diversity of these proteins in synaptic regulation. For instance, the function, if any, of
postsynaptic neurexins in regulating basal glutamatergic neurotransmission remains to be seen.
For this reason, we examined whether acute knockdown of postsynaptic neurexins had any

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functional effect on synaptic regulation within the hippocampus. We find that knockdown of
NRXN1α and NRXN1β does not perturb synaptic function in either CA1 pyramidal or DG
granule neurons of the hippocampus. Further, we find that knockdown of all α- and β-NRXN’s
(NRXN TKD) has no significant effect on synaptic function in either CA1 pyramidal or DG
granule neurons. In Schaffer collateral - CA1 pyramidal synapses, however, NRXN TKD trends
toward decreasing both AMPAR- and NMDAR-mediated eEPSCs – suggesting another neurexin
isoform may contribute to the regulation of these synapses. Altogether, this work contributes to
the growing body of literature detailing the specificity and multifunctionality of various neurexin
isoforms in synaptic regulation.
5.2 Materials & Methods
Experimental Constructs: Previously validated RNAi target sequences against NRXN1 (5’-
GTGCCTTCCTCTATGACAACT-3’), NRXN2 (5’-GAACAAAGACAAAGAGTAT-3’) and
NRXN3 (5’-ATGCTACACTTCAGGTGGACA-3’) were used for the NRXN TKD knockdown
experiments (Zhang et al., 2010; Gokce and Südhof 2013). Each shRNA sequence targets both α
and β isoforms. The same NRXN1-shRNA target sequence was used for the neurexin-1
knockdown experiment. Each shRNA sequence was independently subcloned behind the H1
promoter region of a GFP-expressing pFHUGW expression vector. All plasmids were confirmed
by DNA sequencing.
Electrophysiology: All experiments were performed in accordance with NIH Guidelines
for the Care and Use of Laboratory Animals, and all procedures were approved by the
Institutional Animal Care and Use Committee of the University of Southern California. 400 μm
rat organotypic entorhino-hippocampal slice cultures were prepared from both male and female

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post-natal day 6-8 (P6-8) Sprague Dawley rats as previously described (Stoppini et al., 1991;
Prang et al., 2001; Bonnici and Kapfhammer, 2009; Sadybekov et al., 2017; Tian et al., 2018).
Tissue was isolated and a MX-TS tissue slicer (Siskiyou) was used to make 400 μm transverse
sections. Tissue slices were placed on squares of Biopore Membrane Filter Roll (Millipore) and
placed on Millicell Cell Culture inserts (Millipore) in 35 mm dishes. Slices were fed 1mL of
culture media containing MEM + HEPES (Gibco Cat#12360-038), horse serum (25%), HBSS
(25%) and L-glutamine (1 mM). Media was exchanged every other day. Slices with large portions
of entorhinal cortex were visually identified after slicing. These slices were selected and plated
for use in our experiments, and presence of entorhinal cortex was again confirmed when selecting
slices appropriate for data acquisition. Whole-cell recordings were performed on day in vitro 7-8
(DIV7-8). During recordings, slices were maintained in room-temperature artificial cerebrospinal
fluid (aCSF) containing 119 mM NaCl, 2.5 mM KCl, 1 mM NaH2PO4, 26.2 mM NaHCO3 11 mM
glucose, 4 mM CaCl2, and 4 mM MgSO4. 5 μM 2-chloroadenosine and 0.1 mM picrotoxin were
also added to the aCSF to dampen epileptiform activity and block GABAA receptor activity,
respectively. Osmolarity was adjusted to 310-315 mOsm. aCSF was saturated with 95% O2/5%
CO2 throughout the recording. Borosilicate recording electrodes were filled with an internal
whole-cell recording solution containing 135 mM CsMeSO4, 8 mM NaCl, 10 mM HEPES, 0.3
mM EGTA, 5 mM QX-314, 4 mM Mg-ATP, and 0.3 mM Na-GTP. Osmolarity was adjusted to
290–298 mOsm, and pH-buffered at 7.3–7.4.
DG granule neurons and CA1 pyramidal neurons were identified using differential
interference phase contrast microscopy, while GFP-expressing transfected neurons were
identified using epifluorescence microscopy. Dual whole-cell recordings of either neuronal
subtype were made through simultaneous recordings from a transfected neuron and a

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neighboring, untransfected control neuron. Synaptic responses were evoked by stimulating with a
monopolar glass electrode filled with aCSF in the stratum radiatum for CA1 recordings and the
perforant pathway for DG granule neuron recordings. Membrane holding current, pipette series
resistance, and input resistance were monitored throughout recording sessions. Data were
acquired using a Multiclamp 700B amplifier (Molecular Devices), filtered at 2 kHz, and digitized
at 10 kHz. AMPAR-evoked EPSCS (-eEPSCs) were measured at -70mV. NMDAR-eEPSCs were
measured at +40 mV and were temporally isolated by measuring amplitudes 150 ms following the
stimulus, at which point the AMPAR-eEPSC has completely decayed. Data analysis was
performed using Igor Pro (Wavemetrics). In the scatter plots for simultaneous dual whole-cell
recordings, each open circle represents one paired recording, and the closed circle represents the
average of all paired recordings. No more than one paired recording was performed on any given
entorhino-hippocampal slice.
Biolistic Transfection: Sparse biolistic transfections were performed on DIV1 as
previously described (Stoppini et al., 1991; Schnell et al., 2002; Lu et al., 2009). 50μg of mixed
plasmid DNA was coated on 1μm-diameter gold particles in 0.5 mM spermidine, precipitated
with 0.1 mM CaCl2, and washed four times in pure ethanol. The DNA-coated gold particles were
then coated onto PVC tubing, dried briefly using ultra-pure N2 gas, and stored at 4 °C in
desiccant. Before use, the gold particles were brought up to room temperature and delivered to
slice cultures via a Helios Gene Gun (BioRad). Construct expression was confirmed by GFP or
mCherry epifluorescence.
Experimental Design and Statistical Analyses: All electrophysiological data are
expressed as mean ± standard error measurement (SEM). Statistical significance was determined
using Wilcoxon signed-rank test for paired dual whole-cell patch clamp data. Data were analyzed

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using IGOR Pro (Wavemetrics RRID:SCR_000325) or KaleidaGraph (Synergy Software
RRID:SCR_014980) and graphed using Microsoft Excel (RRID:SCR_016137). All p-values
<0.05 were considered significant and denoted with a single asterisk. All error bars represent
standard error measurement. For all experiments, at least 4 male and female rat pups were used.
Sample sizes in the present study are similar to those reported in the literature (Herring and
Nicoll, 2016; Incontro et al., 2018).
5.3 Results
Knockdown of NRXN1α and NRXN1β (NRXN1 KD) does not perturb basal glutamatergic synapse
function in CA1 pyramidal or DG granule neurons.
Utilizing a previously validated shRNA knockdown sequence for NRXN1, we generated an inhouse vector. This shRNA sequence targets both α and β isoforms (Zhang et al., 2010; Gokce and
Südhof 2013). Following this, we employed a biolistic transfection method to express the
NRXN1-shRNA construct in neurons of rat organotypic entorhino-hippocampal slice cultures
(Elias et al., 2008; Paskus et al., 2019; Tian et al., 2021). These slices provide a method to
examine the functional effects of a genetic modification on hippocampal neurons within their
native circuitry (Stoppini et al., 1991; Schnell et al., 2002). 6 days after transfection, we record
AMPA- and NMDA-receptor evoked excitatory postsynaptic currents (AMPAR- and NMDAReEPSCs) from transfected and neighboring, untransfected control neurons simultaneously during
Schaffer collateral (Fig 5.2A) or perforant pathway stimulation (Fig. 5.2B). This approach allows
for a pairwise, internally controlled comparison of the consequences of our acute genetic
manipulation. We find that knockdown of NRXN1 in CA1 pyramidal neurons produces no
significant change to either AMPAR-eEPSC amplitude (n = 9 pairs, p = 0.91, Wilcoxon signed-

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rank test; Fig. 5.2C, E) or NMDAR-eEPSC amplitude (n = 4 pairs, p = 0.88, Wilcoxon signedrank test; Fig. 5.2F, H). Similarly, we find knockdown of NRXN1 in DG granule neurons results
in no alterations to AMPAR-eEPSC amplitude (n = 6 pairs, p = 0.44, Wilcoxon signed-rank test;
Fig. 5.2D, E) or NMDAR-eEPSC amplitude (n = 5 pairs, p = 1, Wilcoxon signed-rank test; Fig.
5.2G, H). Together, this data suggests neurexin-1 does not have a significant postsynaptic role in
regulating Schaffer collateral-CA1 or perforant pathway-DG granule synapses.
Knockdown of all neurexins (NRXN1, NRXN2, and NRXN3) does not perturb basal
glutamatergic synapse function in CA1 pyramidal or DG granule neurons.
Utilizing previously published shRNA knockdown sequences for NRXN1, NRXN2, and NRXN3,
we generated independent in-house vectors. Each shRNA sequence targets both α and β variants
(Zhang et al., 2010; Gokce and Südhof 2013). The NRXN1-shRNA sequence is identical to the
sequence used above. We find that knockdown of all neurexins (NRXN TKD) in CA1 pyramidal
neurons produces no significant change to either AMPAR-eEPSC amplitude (n = 11 pairs, p =
0.08, Wilcoxon signed-rank test; Fig. 5.3C, E) or NMDAR-eEPSC amplitude (n = 10 pairs, p =
0.08, Wilcoxon signed-rank test; Fig. 5.3F, H). While not statistically significant, knockdown of
all neurexins trends toward reducing both AMPAR and NMDAR currents in Schaffer collateral -
CA1 pyramidal synapses. We also find knockdown of all neurexins at perforant path - DG
granule synapses produces no significant alterations to AMPAR-eEPSC amplitude (n = 7 pairs, p
= 0.58, Wilcoxon signed-rank test; Fig. 5.3D, E) or NMDAR-eEPSC amplitude (n = 8 pairs, p =
0.15, Wilcoxon signed-rank test; Fig. 5.3G, H).

93
5.4 Discussion
Neurexins are a family of presynaptic cell adhesion proteins. Mammals have three
neurexin genes (NRXN1, NRXN2, NRXN3), each of which encode an α and β isoform (Missler
and Südhof, 1998). Immense alternative splicing produces thousands of transcript variants, many
of which show distinct expression patterns in the brain (Treutlein et al., 2014; Fuccillo et al.,
2015). In-situ hybridization shows neurexin mRNA is abundantly expressed within the
hippocampus, though subregion expression varies significantly between isoforms (Uchigashima
et al., 2019).
Most neurexin expression has been detected in presynaptic terminals (Ushkaryov et al.,
1992; Ullrich et al., 1995). Despite this, there is growing evidence that neurexins are found
postsynaptically as well (Peng et al., 2004; Kattenstroth et al., 2004; Taniguchi et al., 2007;
Fairless et al., 2008). The function of postsynaptic neurexins, if any, in synaptic regulation
remains to be seen. One study found cis-expressed β-neurexins inactivate neuroligin-1 and
promote the destabilization of synapses (Taniguchi et al., 2007). Another study found that loss of
α-neurexins induces postsynaptic NMDAR changes in a cell-autonomous manner (Kattenstroth et
al., 2004). Thus, it remains possible that neurexins have a functional role in the postsynaptic
regulation of glutamatergic synapses.
Neurexins contain a protein 4.1 binding domain near their transmembrane region
(Biederer and Südhof, 2001; Reissner et al., 2013). Previous work has shown 4.1N forms a
tripartite complex with CASK and neurexin-1 presynaptically, supporting F-actin nucleation
(Biederer and Südhof, 2001). Due to our previous work detailing a postsynaptic role for 4.1N in
the regulation of perforant path-DG granule synapses, we speculated that this function may
depend on an interaction with neurexin-1. Further, neurexin-1 isoforms show high expression in

94
hippocampal CA1 pyramidal and DG granule neurons (Uchigashima et al., 2019). As such, we
examined the effects of an acute, postsynaptic neurexin-1 knockdown on glutamatergic synapse
function. We find knockdown of neurexin-1 does not alter basal synaptic transmission in either
Schaffer collateral-CA1 or perforant path-DG granule synapses. This work suggests that
neurexin-1 does not have a critical, postsynaptic role in the regulation of these hippocampal
synapses. Alternatively, it remains possible that neurexin-1 has a postsynaptic role in regulating
another synapse type within the brain. Future work may wish to assess the functional effects of an
acute neurexin-1 knockdown in other brain regions with high levels of expression.
To assess whether another neurexin may contribute to the postsynaptic regulation of
hippocampal synapses, we acutely knocked down all neurexins in organotypic hippocampal slice
cultures. We find knockdown of all neurexins does not significantly disrupt glutamatergic
neurotransmission in either Schaffer collateral-CA1 pyramidal or perforant path-DG granule
synapses. Despite this, knockdown of all neurexins in CA1 pyramidal neurons trends towards
reducing both AMPAR- and NMDAR- mediated currents. It remains possible that another
neurexin isoform has a small, partially redundant role in regulating Schaffer collateral- CA1
pyramidal synapses. Going forward, it will be of great interest to assess the functional effects of
loss of individual neurexin isoforms at Schaffer collateral- CA1 synapses.
Altogether, this work contributes to the growing body of literature examining the diverse
roles of neurexins in glutamatergic synapse function. Neurexin isoforms are vast and strongly
vary in their expression patterns, binding affinity, and synaptic functions. While a great deal is
known about the neurexin family, much more remains to be discovered about the diversity of
these cell adhesion proteins.

95
Figure 5.2 Knockdown of neurexin-1 does not disrupt synaptic transmission at CA1
pyramidal or DG granule neuron synapses.

96
(A) Schematic representation of electrophysiological recording setup for CA1 pyramidal
neurons. (B) Schematic representation of electrophysiological recording setup for DG granule
neurons. (C) Knockdown of neurexin-1 in CA1 pyramidal neurons does not significantly affect
AMPAR-eEPSC (n = 9 pairs) or (F) NMDAR-eEPSC amplitude (n = 4 pairs). (D) Knockdown
of neurexin-1 in DG granule neurons does not significantly affect AMPAR-eEPSC (n = 6 pairs)
or (G) NMDAR-eEPSC amplitude (n = 5 pairs). (C, D, F, G) Scatterplots show eEPSC
amplitudes for pairs of untransfected and transfected cells (open circles) with corresponding
mean ±SEM (filled circles). (E, H) Bar graphs show the average AMPAR-eEPSC and NMDAReEPSC amplitudes (±SEM) of neurons expressing the NRXN1-shRNA (orange, maroon)
normalized to their respective control cell average eEPSC amplitudes (black).

97
Figure 5.3 Knockdown of all neurexins does not disrupt synaptic transmission at CA1
pyramidal or DG granule neuron synapses.

98
(A) Schematic representation of electrophysiological recording setup for CA1 pyramidal
neurons. (B) Schematic representation of electrophysiological recording setup for DG granule
neurons. (C) Knockdown of neurexin-1, neurexin-2, and neurexin-3 (neurexin TKD) in CA1
pyramidal neurons does not significantly affect AMPAR-eEPSC (n = 11 pairs) or (F) NMDAReEPSC amplitude (n = 10 pairs). (D) Knockdown of all neurexins in DG granule neurons does
not significantly affect AMPAR-eEPSC (n = 7 pairs) or (G) NMDAR-eEPSC amplitude (n = 8
pairs). (C, D, F, G) Scatterplots show eEPSC amplitudes for pairs of untransfected and
transfected cells (open circles) with corresponding mean ±SEM (filled circles). (E, H) Bar graphs
show the average AMPAR-eEPSC and NMDAR-eEPSC amplitudes (±SEM) of neurons
expressing the NRXNTKD-shRNA (orange, maroon) normalized to their respective control cell
average eEPSC amplitudes (black).

Abstract (if available)

Abstract The molecular mechanisms governing glutamatergic synapse structure and function are vast and complex. Proper synaptic regulation is critical for healthy brain functioning. Historically, most studies examining synaptic function have focused on Schaffer collateral-CA1 synapses and extrapolated findings to be true for all synapses. Recent technological advances have begun to uncover the immense functional, morphological, and molecular differences between synapse subtypes. Despite this, more work needs to be done to truly understand and appreciate the extent of these differences. 4.1N is a protein whose synaptic function has long eluded researchers, despite its binding to multiple synaptic regulatory proteins. Chapter 2 details a novel, site-specific, postsynaptic role for protein 4.1N in regulating perforant path – granule neuron synapses of the dentate gyrus (DG). This role is partially dependent on 4.1N’s FERM domain. Our results strongly suggest that 4.1N’s FERM domain plays a critical role in GluA1-containing AMPAR trafficking to DG granule synapses. Chapter 3 introduces a second role for protein 4.1N in synaptic regulation. Removal of 4.1N’s SAB domain dramatically increases synaptic AMPAR-mediated currents, highlighting the multifunctionality of 4.1N’s role in maintaining stable AMPAR expression at DG granule synapses. Chapter 3 also investigates potential roles for 4.1R and 4.1B, other 4.1 family members with localized hippocampal expression, in synaptic regulation. Overall, my hope is that the work in this dissertation contributes to the growing body of research uncovering the molecular specificity between synapse subtypes. Specifically, this work further confirms perforant pathway synapses are governed by a unique set of molecular regulators.

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Creator Pushkin, Anna Nadya (author)

Core Title Uncovering cell type-specific roles of proteins involved in glutamatergic synapse regulation

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Neuroscience

Degree Conferral Date 2024-05

Publication Date 05/22/2024

Defense Date 04/30/2024

Publisher Los Angeles, California (original), University of Southern California (original), University of Southern California. Libraries (digital)

Tag 4.1,4.1B,4.1N,4.1R,AMPA receptor,CA1 pyramidal neurons,dendritic spines,DG granule neurons,GluA1,glutamatergic synapses,hippocampus,molecular regulators,neurexin,NMDA receptor,OAI-PMH Harvest,synapse,synaptic plasticity

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