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Two Novel Photoactivatable Phosphodiesterases to Independently Hydrolyze cAMP and cGMP
by
Fiona Bergin
A thesis submitted in conformity with the requirements for the degree of Masters of Science Department of Molecular Genetics
University of Toronto
© Copyright by Fiona Bergin 2017
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Two Novel Photoactivatable Phosphodiesterases to
Independently Hydrolyze cAMP and cGMP
Fiona Bergin
Master of Science
Department of Molecular Genetics University of Toronto
2017
Abstract cAMP and cGMP are universal second messengers and have been implicated in synaptic
plasticity underlying learning and memory. Pharmacological and genetic manipulations highlight
the role of postsynaptic cAMP/cGMP in synaptic plasticity; however these techniques alone lack
the precision to understand their rapid and dynamic spatiotemporal roles. To non-invasively
control the rapid molecular activity in living neurons, I developed genetically engineered
photoactivatable PDE4 (PhPDE4) and PDE5 (PhPDE5) to independently degrade cAMP/cGMP
by light. To optimize their light-dependent activities, I validated their single-photon and two-
photon photoactive properties in vitro. These enzymes showed substrate specificity and rapid
millisecond level photoactivation. For in vivo application, I demonstrated the suppression effect
of cAMP in synapse structural plasticity by two-photon photoactivation of PhPDE4. In
combination with bacterial photoactivatable enzymes (PAC, BLGC) to enhance cAMP/cGMP
signaling, these novel light sensitive enzymes will enable us to understand the rapid
spatiotemporal role of cAMP/cGMP in synaptic plasticity and brain function.
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Acknowledgments I would like to first and foremost thank my supervisor Dr. Kenichi Okamoto. Your continued
guidance, support and laughter greatly enriched my graduate studies.
I am grateful for my committee advisors, Dr. Boulianne and Dr. Hui, for their helpful
suggestions and guidance throughout my Master’s degree.
I am thankful for all the members of Dr. Okamoto’s lab, Jelena Borovak, Thomas Luyben,
Megan Valencia and Hang Li for all of your technical guidance and friendly chats!
I sincerely thank my fiancé Garrett Cannella and my family for their constant love, support and
encouragement throughout my studies.
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Table of Contents
Acknowledgments ................................................................................................................ iii
Table of Contents .................................................................................................................. iv
List of Tables ........................................................................................................................ vii
List of Figures ...................................................................................................................... viii
Chapter 1 Introduction ........................................................................................................... 1
Introduction ..................................................................................................................... 1 1
1.1 Overview ..............................................................................................................................1
1.1.1 Scientific method and the need for technological advancements ..........................................1
1.2 Optogenetic tools .................................................................................................................2
1.2.1 Introduction .............................................................................................................................2
1.2.2 Channelrhodopsin: inducing neuronal activity ........................................................................2
1.2.3 Halorhodopsin: suppressing neuronal activity ........................................................................3
1.2.4 Other optogenetic tools: manipulating protein function ........................................................4
1.3 Optogenetic Techniques offer Specific Experimental Control .................................................5
1.3.1 Temporal and Spatial Control of Optogenetic tools ................................................................5
1.3.2 Optogenetics enables manipulation of specific cell types .......................................................7
1.4 Neuroscience ........................................................................................................................8
1.4.1 Synaptic Plasticity .................................................................................................................. 88
1.4.2 Structural Synaptic Plasticity ................................................................................................. 10
1.4.3 LTP and memory ................................................................................................................... 10
1.4.4 Critical intracellular signaling of LTP ..................................................................................... 11
1.5 Optogenetic manipulation of cAMP and cGMP signaling ...................................................... 14
1.5.1 Photoactivatable adenylyl cyclase (PAC) & Blue light sensitive guanylyl cyclase (BLGC) ..... 14
1.5.2 Light Activatable Phosphodiesterase (LAPD) ........................................................................ 14
1.6 Rational and aims of this study ........................................................................................... 16
Chapter 2 Materials and Methods ........................................................................................ 17
Materials and Methods .................................................................................................. 17 2
2.1 Enzyme Design ................................................................................................................... 17
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2.1.1 PhPDE enzyme design ........................................................................................................... 17
2.1.2 Protein expression and purification ...................................................................................... 17
2.2 In Vitro photoactivation assay by ELISA ............................................................................... 18
2.2.1 One photon illumination ....................................................................................................... 18
2.2.2 Two photon illumination ....................................................................................................... 19
2.3 Expression and Validation of PhPDEs in Neurons ................................................................. 19
2.3.1 Slice Culture .......................................................................................................................... 19
2.3.2 Expression in Neurons ........................................................................................................... 20
2.3.3 Structural Long Term Potentiation (sLTP) experiments ........................................................ 20
2.4 Calculations and Statistical Analysis .................................................................................... 21
Chapter 3 Photoactivatable Phosphodiesterase 4 (PhPDE4) .................................................. 22
Photoactivatable Phosphodiesterase 4 (PhPDE4) ............................................................ 22 3
3.1 Introduction ....................................................................................................................... 22
3.2 Results ............................................................................................................................... 24
3.2.1 Design of Photoactivatable Phosphodiesterase 4 (PhPDE4) ................................................. 24
3.2.2 Optimization of the PhPDE4 light dependent photoactivation ............................................ 25
3.2.3 Characterization of PhPDE4 using one-photon light (LED) ................................................... 26
3.2.4 Two-photon characterization of PhPDE4 .............................................................................. 29
3.2.5 Establish soluble and membrane bound forms of PhPDE4................................................... 31
3.2.6 Validation of PhPDE4 activity in living neurons .................................................................... 33
3.2.7 Photoactivation of PhPDE4 for future mouse behavioral experiments ................................ 36
3.3 Summary of Chapter 4 ........................................................................................................ 37
Chapter 4 Photoactivatable Phosphodiesterase 5 (PhPDE5) .................................................. 38
Photoactivatable Phosphodiesterase 5 (PhPDE5) ............................................................ 38 4
4.1 Introduction ....................................................................................................................... 38
4.2 Results ............................................................................................................................... 40
4.2.1 PhPDE5 enzyme design ......................................................................................................... 40
4.2.2 Optimization of the PhPDE5 light dependent photoactivation ............................................ 41
4.2.3 Characterization of PhPDE5 using one-photon light (LED) ................................................... 42
4.2.4 Two-photon characterization of PhPDE5 .............................................................................. 45
4.2.5 Establish soluble and membrane bound forms of PhPDE5................................................... 47
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4.2.6 Illumination technique of PhPDE5 for future mouse behavioral experiments ..................... 49
4.3 Summary of Chapter 4 ........................................................................................................ 49
Chapter 5 Comparison of photo activity between PhPDEs and LAPD ..................................... 50
Comparison of photo activity between PhPDEs and LAPD ............................................... 50 5
5.1 Introduction ....................................................................................................................... 50
5.2 Results ............................................................................................................................... 51
5.2.1 Comparison of the light sensitive activation between LAPD, PhPDE4 and PhPDE5 ............. 51
5.2.2 Two-photon activation of LAPD ............................................................................................ 52
5.2.3 Photoactivation properties of purified LAPD, PhPDE4 and PhPDE5 ..................................... 53
5.2.4 Establish soluble and membrane bound forms of LAPD ....................................................... 53
5.2.5 Preparation of LAPD lenti viral construct for in vivo applications ........................................ 55
5.3 Summary of Chapter 5 ........................................................................................................ 56
Chapter 6 Discussion ............................................................................................................ 57
Discussion ...................................................................................................................... 57 6
6.1 Summary of the project ...................................................................................................... 57
6.2 Photoactive properties of PhPDEs ....................................................................................... 58
6.3 In vivo validation of PhPDEs in living neurons ...................................................................... 60
6.4 Comparisons and limitations ............................................................................................... 60
6.5 Future Applications ............................................................................................................. 62
6.6 Conclusion .......................................................................................................................... 62
References ........................................................................................................................... 63
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List of Tables Table 1 Photoactivation of purified LAPD, PhPDE4 and PhPDE5 using one-photon and two-photon
excitation light…………………………………………………………………………………….... 53
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List of Figures
Chapter 1
Figure 1-1 One-Photon and two-photon excitation properties. …………………..…………………..….6
Figure 1-2 Two-photon laser excitation spatial resolution………………………..…………………..….7
Figure 1-3 The hippocampus is a critical structure for long term memory formation…….…………..….9
Figure 1-4 cAMP and cGMP postsynaptic signaling……………………………………………………13
Figure 1-5 Current optogenetic tools to manipulate cAMP and cGMP signaling with light……………15
Chapter 3
Figure 3-1 Protein sequence alignment of the C-terminal catalytic domains in PDE2A used for LAPD and PDE4B. ……………………………………………………………………………………..………...25
Figure 3-2 Design of photoactivatable PDE4 variants and their activity………………………..…..…....26
Figure 3-3 cAMP dependent photoactivity of PhPDE4 has a rapid on and off response in vitro.….……27
Figure 3-4 PhPDE4 is activated by various light intensities and wavelengths using one-photon light......28
Figure 3-5 PhPDE4 is more efficiently activated by longer two-photon excitation and shows rapid and light intensity dependent activation at 1,100 nm…………………………………………………...……..30
Figure 3-6 In vitro one-photon photoactivation between membrane bound and soluble forms of PhPDE4……..……………………………………………………………………………………..………31
Figure 3-7 Subcellular localization of membrane bound and soluble forms of PhPDE4 in living neurons……..……………………………………………………………………………….……..………32
Figure 3-8. PhPDE4 activation at single dendritic spines suppresses cAMP effect during structural LTP……..………………………………………………………………………………...………..………34
Figure 3-9 PhPDE4 activation at single dendritic spine causes little change during sLTP induction but significantly suppresses the cAMP dependent potentiation of sLTP.………..……………………………35
Figure 3-10 In vitro photoactivation of PhPDE4 using the implantable fiber optic LED light source for in vivo mouse experiments. ……..…………….……………………………………………………..………36
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Chapter 4
Figure 4-1 Protein sequence alignment of the C-terminal catalytic domains in PDE2A used for LAPD and PDE5A. ……..………………………………………………………………………………..………41
Figure 4-2 Design of photoactivatable PDE5 variants and their activity.………………………..………42
Figure 4-3 cGMP dependent photoactivity of PhPDE5 has a rapid on and off response in vitro………..43
Figure 4-4 PhPDE5 is activated by various light intensities and wavelengths using one-photon light.……..………………………………………………………………………………………..…….…44
Figure 4-5 PhPDE5 is more efficiently activated by longer two-photon excitation and shows rapid and light intensity dependent activation at 1,100 nm………………………………………………………….46
Figure 4-6 In vitro one-photon photoactivation between membrane bound and soluble forms of PhPDE5. ……..…………………………………………………………………………………..…………….……47
Figure 4-7 Subcellular localization of membrane bound and soluble forms of PhPDE5 in living neurons. ……..…………………………………………………………………………………..………………….48
Figure 4-8 In vitro photoactivation of PhPDE5 using the implantable fiber optic LED light source for in vivo mouse experiments. ..………………………………………………………………………………..49
Chapter 5
Figure 5-1 In vitro photoactivation of PhPDE4, PhPDE5 and LAPD on cAMP and cGMP levels. ……51
Figure 5-2 Two-photon excitation efficiently induces the degradation of both cAMP and GMP ..……..52
Figure 5-3 Subcellular localization of membrane bound and soluble forms of LAPD in living neurons 54
Figure 5-4 Validate expression and light dependent activity of pLenti-LAPD in HEK293 cells….…..…55
Chapter 6
Figure 6-1 Schematic diagram of the photocycle of PhPDEs……………………………………………58
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List of Abbreviations AC Adenylyl Cyclase
BLGC Blue Light Sensitive Guanylyl Cyclase
bPAC Bacterial Photoactivatable Adenylyl Cyclase
CA1 Cornu Ammon1 (type of Hippocampal Pyramidal Neuron)
CaMKII Calcium calmodulin-dependent protein kinase II
cAMP Cyclic adenosine monophosphate
cGMP Cyclic guanosine monophosphate
ChR2 Channelrhodopsin 2
CRE CREB responsive element
Cre-LoxP Cre recombinase – Locus of X-over P1
CREB cAMP response element binding protein
DISC1 Disrupted in schizophrenia
E-LTP Early Long Term Potentiation
ELISA Enzyme Linked Immunosorbent Assay
GAP43 Growth Associated Protein – 43
GC Guanylyl Cyclase
GFP Green Fluorescent Protein
HEK293 Human Embryonic Kidney 293 cells
L-LTP Late Long Term Potentiation
LAPD Light activatable phosphodiesterase
LED Light emitting diode
LTD Long Term Depression
LTP Long Term Potentiation
MLS Membrane localization sequence
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NA Numerical aperture
NMDAR N-methyl-D-aspartate- type glutamate receptor
NO Nitric Oxide
NOS Nitric Oxide Synthase
NpHR Halorhodopsin
PAS-GAF-PHY Per-arnt-sim - cGMPPDE/adenylyl cyclase/Fh1A - Phytochrome domain
PDE Phosphodiesterase
PhPDE4 Photoactivatable Phosphodiesterase 4
PhPDE5 Photoactivatable Phosphodiesterase 5
PKA Protein Kinase A
PKG Protein Kinase G
RFP Red Fluorescent Protein
sLTP Structural Long Term Potentiation
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Chapter 1 Introduction
Introduction
1.1 Overview
1.1.1 Scientific method and the need for technological advancements
To understand molecular mechanisms of learning and memory, pharmacological and genetic
manipulations of various molecules have been used in the brain. However, those traditional
approaches are irreversible and limited in their spatial and temporal manipulation (Dugue,
Akemann, & Knopfel, 2012). Pharmacological approach using specifically designed chemical
inhibitors or stimulators enables perturbations of signalling pathways at the molecular level. This
approach highlights the link between different molecules and their cellular function. However,
pharmacological manipulation is limited by its spatial specificity as administration in vivo causes
widespread cellular changes in the brain. Furthermore, depending on the chemical inhibitor or
stimulator as well as the mode of injection, pharmacological approaches are typically required to
manipulate molecular activity at the minutes to hours’ time scale (Toettcher, Voigt, Weiner, &
Lim, 2011). Another technique widely used is genetic manipulations. Genetic targeting and
transgenic approaches enable precise manipulation of single genes. Knocking out or
overexpressing single genes in vivo enables observation of associated behavioural changes.
While this technique has great molecular specificity, it lacks the ability to manipulate synapse
level function. Also, to control the temporal alteration of genetic manipulation is available using
inducible systems however this takes up to a few hours (Weber et al., 2012).
For manipulating with precise temporal control as well as spatial specificity including
subcellular and cellular targeting I established an optogenetic approach to suppress cAMP and
cGMP for studying their functions in the brain. This optogenetic manipulation is reversible with
rapid onset and offset with minimal invasion. In combination with pharmacological and genetic
techniques, this approach will be a critical advanced technique for neuroscience research.
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1.2 Optogenetic tools
1.2.1 Introduction
Optogenetics is defined as “the combination of genetic and optical methods to achieve
gain or loss of function of well-defined events in specific cells of living tissue” (Deisseroth,
2011). Various types of photoreceptors are found in nature and these light sensitive proteins
contain different chromophores, which can absorb specific wavelengths of light from ultra
violet-B to near-infrared light (Rockwell et al., 2014). Typically, the absorption of light by the
chromophore in the light sensitive domain induces a conformational change resulting in altered
function of the receptor (Ziegler & Moglich, 2015). The light sensitive activity causes light-
sensitive locomotion of organisms to move towards or away from light or functions to initiate a
biochemical cascade for energy production (Jenkins, 2017; Ueki et al., 2016). To apply this
property to neuroscience research these light sensitive molecules were characterized and
optimized for controlling neuron activity by light.
1.2.2 Channelrhodopsin: inducing neuronal activity
Channelrhodopsin (ChR2), a single component light activated cation channel was discovered
from algae Chlamydomonas reinhardtii (Nagel et al., 2002). This channel absorbs blue light,
causing a conformational change, opening the channel and enabling the passive diffusion of
cation ions down its concentration gradient. This channel enables Chlamydomonas reinhardtii to
find optimal conditions for photosynthesis as it stimulates swimming towards light in dim
conditions or moving away from excess amount of light (Erickson, Wakao, & Niyogi, 2015;
Nagel et al., 2002; Nagel et al., 2003). When it is expressed in living neurons, illumination with
blue light causes the channels to open within a millisecond, inducing rapid depolarization of the
membrane potential, which induces action potentials and thus neuronal excitability (Boyden,
Zhang, Bamberg, Nagel, & Deisseroth, 2005). This discovery accelerated the search for other
photoreceptors for the optogenetic application to control neuronal activity.
ChR2 was used to examine the properties of memory storage. Current knowledge in this
area suggests that during a distinct event, strong simultaneous activation of a discrete subset of
neurons creates a specific memory trace. Reactivation of this specific trace is believed to
stimulate memory recall and should be distinct and independent of other memories. Liu and
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colleagues specifically expressed ChR2 only in the neurons activated during a fear memory (Liu
et al., 2012). Illumination of ChR2 expressed only in this specific neural circuit, in a neutral
context elicited the same fear memory recall. Verifying that specific memories are stored in
discrete neural circuits. This experiment highlights the application of optogenetic tools for
neuroscience research.
1.2.3 Halorhodopsin: suppressing neuronal activity
A perfect tool to complement such experimental manipulation would be a light sensitive
channel that can silence neural activity with the same precision. The discovery of halorhodopsin,
a chloride pump from Natronomonas pharasonis (NpHR) provided this complementary tool for
ChR2, to turn neural activity off (Kolbe, Besir, Essen, & Oesterhelt, 2000). When expressed in
living neurons, NpHR protein was found to hyperpolarize the membrane potentials and silence
neural activity upon illumination within millisecond precision. Interestingly, NpHR is optimally
activated by yellow light instead of blue light used for ChR2. It was demonstrated that, ChR2
and NpHR can be co-expressed and independently activated using different wavelengths of light
such that blue light induces activity of neurons and yellow light suppresses activity (F. Zhang et
al., 2007).
These optogenetic tools work in vivo as demonstrated in Caenorhabditis elegans. The
illumination of blue and yellow light induced the activation and silencing of motor neuron
activity respectively in C. elegans (F. Zhang et al., 2007). Furthermore, the activation of ChR2 in
muscle cells of C. elegans caused an instant increase in swimming motion whereas activation of
NpHR by yellow light instantly blocked this movement. Together these optogenetic tools enable
scientists to study the effect of activating or silencing neuronal activity, within a biologically
relevant time scale to study the functions of specific neural circuits.
These optogenetic tools were also applied to address brain diseases such as the neural connection
to anxiety disorders, which encompass numerous psychiatric diseases such as posttraumatic
stress disorder, generalized anxiety disorder and panic disorder (Tye et al., 2011) (Lieb, 2005).
In a rodent anxiety model, specific stimulation of ChR2 in a discrete population of excitatory
neurons in the amygdala showed significantly reduced anxiety like behaviour. Furthermore,
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selective illumination and therefore inhibition of this same projection using NpHR to
hyperpolarize this pathway resulted in anxiety. This experiment was repeated on a different
neural projection in the brain composed of the same cell types but showed no effect on anxiety
levels when ChR2 or NpHR were activated (Tye et al., 2011). Therefore, using this optogenetic
approach a specific cellular pathway rather than cell type was isolated to be important for
anxiety. Thus, optogenetic approach offers superior specific experimental control and will have a
big impact on basic neuroscience research as well as understanding diseased states.
1.2.4 Other optogenetic tools: manipulating protein function
The field of optogenetics accelerated as optogenetic tools moved from manipulating whole cell
activities to perturbing intracellular molecular functions (Guglielmi, Falk, & De Renzis, 2016).
Critical cellular activities are regulated by kinase and phosphatase activity. Currently there is a
light sensitive Raf kinase, which induces reversible activity of the Raf/MEK/ERK pathway upon
illumination (Krishnamurthy et al., 2016). Conversely, there is a light sensitive
phosphatidylinositol 5-phosphatase, which is recruited to the cell membrane upon illumination
and causes rapid dephosphorylation of its downstream targets (Idevall-Hagren, Dickson, Hille,
Toomre, & De Camilli, 2012). Some other optogenetic enzymes recently developed include
Photoactivatable Rac1, which regulates actin cytoskeleton dynamics (Wu et al., 2009) or a light
inducible protein oligomerization (CRY2olig), which can be used to examine protein interactions
in living cells (Taslimi et al., 2014). All of these engineered optogenetic enzymes now enable the
perturbation of various intracellular activities with subcellular resolution and fast, seconds order
precision (Guglielmi et al., 2016).
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1.3 Optogenetic Techniques offer Specific Experimental Control These light sensitive tools are superior manipulators of biological activity by their activation
speed, spatial control and the ability to restrict their expression to specific neuronal populations
(Deisseroth, 2011).
1.3.1 Temporal and Spatial Control of Optogenetic tools
Optogenetic tools offer millisecond control of excitation and inhibition of neural activity in
specific populations. Both ChR2 and NpHR have confirmed millisecond activation and silencing
of living neurons. This light dependent response advances the study of rapid temporal functions
of specific neuronal populations in the brain functions and related diseases (F. Zhang, Wang,
Boyden, & Deisseroth, 2006).
Another benefit of optogenetics is spatial control of neuronal activation by light. For example,
one photon illumination can be used to activate specific brain region or neural circuit expressing
light sensitive proteins to understand their roles in the brain functions and related diseases.
Moreover two-photon laser excitation with a fine focal point (~1μm3) using a high numerical
aperture (NA) objective lens, can be employed to localize activation to subcellular structures
such as single synapses to understand the impact on the function of microcircuits with synapse
level precision (Boyden et al., 2005).
Maria Goppert-Mayer first theorized the concept of two-photon absorption in 1931. One-
photon excitation of a fluorophore occurs when fluorophores are excited from their electronic
ground state to an excited state. Maria believed that this same excitation of a fluorophore could
be achieved by the simultaneous absorption of two less energetic photons from the infrared
spectra range (Figure 1-1). After this effect was validated in 1963, two-photon microscopy was
established in the 1990s (Denk, Strickler, & Webb, 1990; Kaiser & Garrett, 1961). To increase
the probability of simultaneous absorption of two photons, a femtosecond pulse laser is used for
two-photon microscopy. This two-photon excitation uses longer wavelengths of infrared light
therefore enabling deeper tissue penetration and spatially restricts activation to only the focal
point where two photons of light meet (~1μm3) (Figure 1-2). These properties make two-photon
laser microscopy a valuable tool for restricting optogenetic stimulation to a subcellular level.
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Figure 1-1 One-Photon and two-photon excitation properties. (A) One-photon excitation
using the visible light spectrum produces a larger area of illumination compared to (C) Two-
photon excitation focal volume (1µm3) using infrared light (radius = 0.61 x wavelength / NA x
1.414) (B) Excitation occurs when one photon absorption causes excitation between the ground
and excited state. (D) Two photon excitation occurs through the simultaneous absorption of two
lower energy photons (infrared range) if their combined energy is greater than the energy
difference between ground and excited state.
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Figure 1-2 Two-photon laser excitation spatial resolution.
(A) Due to the use of two-photons of light from longer infrared wavelengths of light, two-photon
imaging of mouse brain produces less scattering than confocal imaging resulting in deeper tissue
penetration (~ a few mm). (B) Two-photon microscopy has optimal spatial resolution for
resolving or manipulating single synaptic structures such as the post synaptic dendritic spines
highlighted above by white line.
1.3.2 Optogenetics enables manipulation of specific cell types
In combination with specific promoters, genetically engineered optogenetic tools can be
expressed in specified neuronal populations in the brain. For example, lentivirus packaged with a
CaMKIIα promoter, limits the expression in excitatory neurons and maintains the expression of
the optogenetic tool for more than 6 months in the mouse brain (Wang, Zhang, Szábo, & Sun,
2013).
Cre-LoxP expression system also enables us to limit the expression of optogenetic tools
in specific cells. This system utilizes a transgenic mouse line containing Cre recombinase under
the expression of a cell type specific promoter and either viral vector or mouse line containing
the inverted light sensitive protein flanked by loxP sites. Injection of the virus into these
transgenic mice restricts expression of optogenetic tools to cells expressing Cre recombinase.
This approach has been successful using lentivirus or adeno- associated virus (AAV) to target
optogenetic tools to specific neural cell types (Zeng & Madisen, 2012).
Another targeting method is the neuronal activity-dependent expression of optogenetic
enzymes. The expression of immediate early genes such as c-fos can be used to identify the
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neurons, which were recently activated during a specific learning and behavioral task. These
immediate early genes are transcribed within minutes and associated with neural activity
(Guzowski, 2002). Using the activity-dependent inducible system, optogenetic protein such as
ChR2 can be expressed only in the neurons activated during a particular task. This expression
system requires expression of ChR2 to be driven by the tetracycline-responsive element (TRE)
and a transgenic mouse line where the tetracycline transactivator (tTA) is driven by c-fos
promoter (Ramirez, Tonegawa, & Liu, 2013). Therefore, neural activity induces expression of
immediate early genes such as c-fos driving the expression of tTA to activate ChR2 expression in
only those neurons. Now only the cells activated during that particular training contain the
expressed ChR2 permitting the activation of only that neural circuit using light (Ramirez et al.,
2013).
1.4 Neuroscience
1.4.1 Synaptic Plasticity
Ramon y Cajal first demonstrated that brain tissue is composed of a network of signaling units
called neurons, and these neurons are arranged in functional groups that connect to one another
in a precise circuit. He postulated that memory storage relies on changes in the strength of
connections between activated neurons (Ramón y Cajal, 1995). The activity dependent changes
occur at the synapse, the junction where the presynaptic bouton meets the postsynaptic structure,
called the dendritic spine of another neuron. In the 1940s Donald Hebb supported this hypothesis
coining the phrase “neurons that wire together fire together, neurons that wire apart, fire apart”
(Morris, 1999). To confirm this hypothesis, many researchers looked into the mechanism
underlying how the synapse is strengthened leading to the model of long-term potentiation
(LTP). In 1972 Timothy Bliss and Terje Lomo applied brief high frequency trains of action
potentials in rabbit hippocampal slices resulting in stronger synaptic transmission and the
observation of LTP (Bliss & Lomo, 1973). Long term depression (LTD) was also found in
synapses that undergo prolonged low-frequency stimulation therefore suppressing synaptic
activity (Lynch, Dunwiddie, & Gribkoff, 1977).
The hippocampus is a critical region in the brain enabling long-term memory formation of
declarative memories. There are two different forms of synaptic plasticity in the hippocampus,
post-synaptic dependent and pre-synaptic dependent which occur in different hippocampal neural
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circuits. The induction of post-synaptic dependent LTP requires activation of N-methyl-D-
aspartate (NMDA)- type glutamate receptor (Collingridge, Kehl, & McLennan, 1983). In order
for NMDA receptors to be activated, they need the pairing of glutamate release from the pre-
synaptic terminal and depolarization of the postsynaptic membrane, enabling an influx of
calcium into the postsynaptic spine. This type of LTP is observed in two of the three-
hippocampal pathways (Shaffer collaterals, performant path) initially described by Ramon Y
Cajal using the Golgi staining method (Figure 1-3). The third pathway is non-NMDA receptor
dependent where the mossy fibers synapse onto CA3 neurons and rely on presynaptic dependent
mechanisms for the induction of LTP. Importantly LTP was blocked when calcium signalling
was inhibited by EGTA and conversely, LTP was induced when calcium was added to the
presynaptic cell. Therefore indicating that presynaptic increase in intracellular calcium ions is
critical for LTP induction.
Figure 1-3 The hippocampus is a critical structure for long term memory formation.
(A) The location of the hippocampus in the brain. (B) Drawing of the neural circuitry of the
rodent hippocampus adapted from Ramon y Cajal (1911).
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1.4.2 Structural Synaptic Plasticity
Along with these functional changes structural alterations also occur during synaptic plasticity
and requires reorganization of the major postsynaptic cytoskeletal protein actin. Actin is
regulated by the binding and release of an abundant postsynaptic protein CaMKII (Calcium
calmodulin-dependent protein kinase II) suggesting its importance in regulating structural and
functional plasticity. Our lab found that the postsynaptic CaMKIIβ/actin regulates both
functional and structural synaptic plasticity (Kim et al., 2015; Okamoto, Nagai, Miyawaki, &
Hayashi, 2004).
1.4.3 LTP and memory
Pharmacological inhibition of NMDA receptors blocked LTP and also impaired spatial memory
in the mouse hippocampus using the morris water maze (Morris, Anderson, Lynch, & Baudry,
1986). These early findings highlight the link between LTP and learning and memory as well as
introduced critical intracellular signalling underlying LTP.
Interestingly, LTP shares a similar time scale as memory formation further highlighting the link
between LTP and memory. Early phase of LTP (E-LTP) is typically induced by weak
stimulation, a single train of action potentials, causing an increase in intracellular calcium
leading to short-term structural and functional potentiation. This plasticity enhancement lasts for
up to 3 hours and has been linked to short-term memory formation (Kandel, 2012). Late phase of
LTP (L-LTP) is induced by multiple trains of synaptic stimulation, and causes a more persistent
structural and function plasticity enhancement lasting over 3 hours to months. This plasticity
enhancement is due to stronger stimulation inducing a greater calcium influx leading to trigger
new protein synthesis pathway critical for long-term memory formation and consolidation (Abel
et al., 1997; Frey, Huang, & Kandel, 1993). Application of protein synthesis inhibitors disrupt
long-term memory formation but did not affect short-term memory formation, suggesting a
critical link between L-LTP and long term memory (Frey, Krug, Reymann, & Matthies, 1988).
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1.4.4 Critical intracellular signaling of LTP
cAMP
Earl Sutherland was awarded the Nobel Prize in Physiology and Medicine in 1971 for
discovering cyclic adenosine monophosphate (cAMP). He determined that cAMP was a critical
second messenger that amplifies signal transduction across the cellular membrane (Sutherland,
1972). Neurotransmitters such as serotonin increase cAMP concentration in the brain linking
cAMP signalling and neural activity (Brunelli, Castellucci, & Kandel, 1976). Cyclic AMP
signalling is now known to mediate many activities in the brain including developmental
properties, synaptic plasticity and learning and memory. Using the fruit fly, Drosophila
melanogaster, mutations to the dunce gene, a cAMP-phosphodiesterase (PDE) which degrades
cAMP, caused an increase in cAMP signalling and strong learning deficits (Dudai, Jan, Byers,
Quinn, & Benzer, 1976). However, flies with a mutant adenylyl cyclase which synthesizes
cAMP, rutabaga, resulted in decreased cAMP signalling and defective learning (Levin et al.,
1992). These results suggest a critical link between regulated cAMP signalling and learning and
memory functions. At the cellular level, Eric Kandel, found that short-term memory depicted in
one trial of the gill-withdrawal reflex in the marine slug Aplysia, produced cAMP, which
transiently enhanced neurotransmitter release (Kandel, 2001). Furthermore, multiple training
trials caused long-term memory of the gill-withdrawal reflex lasting days to weeks which
required a more persistent increase in cAMP signalling causing cAMP-dependent protein
synthesis of plasticity related genes critical for long term memory formation. This simple reflex
system demonstrated in the very large (up to 1mm) neuron of Aplysia demonstrated the
important link between cAMP signalling, protein synthesis and synaptic plasticity. The canonical
cAMP pathway involves the hormone signalling, which activates adenylyl cyclase (AC) to
produce cAMP. cAMP then activates protein kinase A (PKA) which translocates into the nucleus
and phosphorylates CREB which binds the CREB responsive element (CRE) leading to protein
synthesis (Kandel, 2012).
12
cGMP
3’5’ cyclic guanosine monophosphate (cGMP) is also a crucial second messenger to regulate
synaptic plasticity. cGMP is synthesized by nitric oxide (NO) stimulation of guanylyl cyclase
(GC), which converts GTP to cGMP. The canonical cGMP-signaling pathway, widely studied in
the CA1 neurons of the Schaffer Collateral pathway of the hippocampus during LTP induction,
includes NMDA receptor mediated Ca2+ influx which stimulates Ca2+/calmodulin-activated
neuronal NO synthase (nNOS) causing NO production in the postsynaptic structure. NO acts as a
retrograde messenger and translocates into the presynaptic structure where it activates GC to
produce cGMP and initiates the activation of cGMP dependent protein kinases (PKG). These
kinases initiate the increase of neurotransmitter release aiding in the presynaptic-dependent
potentiation of the synapse. NO also stimulates postsynaptic GC to activate postsynaptic PKG
for enhancing protein synthesis pathways. Thus, both cAMP and cGMP intracellular signalling
are known to play critical roles in synaptic plasticity.
PDEs
Various neurodegenerative diseases such as Alzheimer’s cause dramatic cognitive deficits for
which treatment is limited. Drug development for these diseases has moved from its original
focus of directly targeting neurotransmitters such as acetylcholine, glutamate and dopamine, to
the downstream signalling pathways in long-term memory formation. One of the major drug
targets for neurodegenerative diseases are phosphodiesterases (PDEs). PDE inhibitors, such as
rolipram, sildenafil and tadalafil have been shown to improve cognitive functions in various
mouse models of Alzheimer’s disease (Garcia-Osta, Cuadrado-Tejedor, Garcia-Barroso,
Oyarzabal, & Franco, 2012). However, the molecular mechanisms are still elusive.
The signalling dynamics between cAMP and cGMP during synaptic plasticity is also still
elusive. One member of the PDE family, PDE2, degrades both cAMP and cGMP. However the
rate at which each cyclic nucleotide is degraded depends on the cellular concentration of each.
Specifically, if there is low concentration of cGMP, then it will bind the N-terminal regulatory
domain and heighten the hydrolysis of cAMP. But if there is a low concentration of cAMP, it can
bind the N-terminal regulatory domain and cause an increase in cGMP hydrolysis. This activity
enables one cyclic nucleotide to be the dominant messenger molecule within that subcellular
micro domain. However, when both cyclic nucleotides are present in high concentrations, they
13
compete for the binding site (Zhao, Greenstein, & Winslow, 2016). This causes important spatial
regulation of cyclic nucleotide signalling as well as temporal regulation by maintaining or
supressing cAMP or cGMP signalling depending on the cellular environment. This interesting
dynamic suggest an important crosstalk mechanism between these cyclic nucleotides however
the functions for such interaction at the cellular and subcellular level is unknown. Due to the
high expression of PDE2 in limbic structures such as the cortex, amygdala and hippocampus,
which are all critical sites for memory formation, maintaining the dynamic signalling interplay
between cAMP and cGMP signalling could be important for synaptic plasticity underlying
learning and memory. Inhibition of PDE2, maintaining both cAMP and cGMP signalling
enhances LTP and showed improvement on various memory tasks (Boess et al., 2004). Two
hours of acute tryptophan depletion, a dietary precursor for serotonin, causes short-term memory
impairment observed using the object recognition test. However using BAY 60-7550, a PDE2
inhibitor, 30 minutes of application before the test improved the short-term memory of these
mice (van Donkelaar et al., 2008). While research using PDE inhibitors has greatly expanded our
knowledge surrounding cAMP and cGMP signalling their spatiotemporal role and crosstalk
interactions during synaptic plasticity are less understood.
Figure 1-4 cAMP and cGMP postsynaptic signalling. Summary of the post-synaptic cAMP
and GMP signalling and the central role of various PDEs.
14
1.5 Optogenetic manipulation of cAMP and cGMP signaling
1.5.1 Photoactivatable adenylyl cyclase (PAC) & Blue light sensitive guanylyl cyclase (BLGC)
Due to the widespread role of cAMP signaling, it is not surprising that a photoactivatable
adenylyl cyclase was found in a bacterium Beggiatoa (Stierl et al., 2011). This single component
blue light sensitive adenylyl cyclase (bPAC) has little activity levels in the dark but rapidly
produces cAMP upon illumination. This optogenetic tool has confirmed light dependent activity
and expresses in hippocampal pyramidal neurons, generating an efficient light sensitive approach
to manipulate cAMP spatiotemporal dynamics during synaptic plasticity. Using this bPAC,
mutagenesis of various residues involved in substrate binding sites, enabled the binding of GTP
instead of ATP and therefore produced cGMP upon illumination with blue light called blue light
sensitive guanylyl cyclase (BLGC). These tools now enable the rapid, reversible and non-
invasive stimulation of cAMP and cGMP signaling independently. To complete this optogenetic
toolkit, I prepared new tools, which independently degrade cAMP or cGMP.
1.5.2 Light Activatable Phosphodiesterase (LAPD)
A new optogenetic enzyme was recently developed, Light Activated Phosphodiesterase (LAPD),
which degrades both cAMP and cGMP upon illumination (Gasser et al., 2014). This enzyme is
composed of the light sensitive phytochrome domain from Deinococcus radiodurans fused to the
catalytic domain of PDE2, which degrades both cAMP and cGMP. The N-terminal regulatory
domain of PDEs differs across variants and contains various sites for the binding of modulators
to regulate their activity or localization properties. The C-terminal domain contains the catalytic
domain where binding and hydrolysis of cyclic nucleotides occur. Major similarities were
observed between the bacterial phytochrome domain of Deinococcus radiodurans and the N-
terminal regulatory GAF domains of PDE2. The structural similarities between these regulatory
domains enabled the replacement of the PDE2 N-terminal regulatory domain with the bacterial
phytochrome domain. Successfully enabling the degradation of both cAMP and cGMP to be
controlled by illumination with blue (455 nm), red (690 nm) and even broadband white light
(Gasser et al., 2014). The development of LAPD which degrades both cyclic nucleotides
provides a promising framework for the development of two novel optogenetic enzymes to
independently degrade cAMP or cGMP enabling the detailed understanding of activating or
blocking these second messengers during synaptic plasticity.
15
Figure 1-5 Current optogenetic tools to manipulate cAMP and cGMP signaling with light.
(A) The current optogenetic toolkit is composed of photoactivatable adenylyl cyclase (PAC) to
produce cAMP, a blue light sensitive guanylyl cyclase that produces cGMP and a light
activatable phosphodiesterase LAPD (PDE2), which degrades both cAMP and cGMP. (B) Venn
diagram dividing the different families of PDEs by their substrate specificity. The red boxes
highlight a phosphodiesterase from each group, which will be used to generate the final
optogenetic tools to specifically degrade cAMP (PDE4) or cGMP (PDE5) independently using
the same strategy employed for LAPD.
16
1.6 Rational and aims of this study In our lab we have established two-photon optogenetic microscopy techniques to light
dependently activate specific enzymes such as bPAC and BLGC at a single synapse resolution in
living hippocampal tissues to produce cAMP and cGMP. To compliment these tools I will create
two new optogenetic enzymes to light dependently degrade cAMP or cGMP. There are 11
families of PDEs encoded by 21 genes, each with unique tissue expressions (Bender & Beavo,
2006). Mammalian PDEs are divided into three types: PDEs that hydrolyze cAMP and cGMP
(PDE1, 2, 3, 10, 11), cAMP specific (PDE4, 7, 8) and cGMP specific (PDE5, 6, 9). By replacing
the PDE2 catalytic domain from LAPD with a cAMP-specific PDE (PDE4) catalytic domain or
cGMP-specific PDE (PDE5) catalytic domain, I prepared novel optogenetic enzymes to suppress
endogenous cAMP and cGMP signaling independently at the single synapse level. These
complementary tools will enable us to study their roles in synaptic plasticity by producing cAMP
or cGMP, or degrading cAMP or cGMP with spatial and temporal control of light.
Goal: To create and validate novel optogenetic enzymes to degrade cAMP and cGMP
independently enabling the spatiotemporal control of intracellular signaling processes mediated
by these second messengers.
Aim1: Develop and validate optogenetic tools to independently degrade cAMP or cGMP upon
illumination
Aim2: Validate and characterize two-photon photoactivation of PhPDE4 and PhPDE5 in vitro
Aim3: Demonstrate the functionality of PhPDEs in living neurons
17
Chapter 2 Materials and Methods
Materials and Methods 2
2.1 Enzyme Design
2.1.1 PhPDE enzyme design
The DNA encoding LAPD (Gasser et al., 2014) was synthesized with codon usage optimized for
Homo sapiens (GenScript, New Jersey, USA). The gene encoding the catalytic domain of Homo
sapiens cAMP-specific phosphodiesterase 4 (HsPDE4B; UniProt PDE4B_HUMAN: amino acids
630-1971) was synthesized (GenScript, NJ, USA) and cGMP-specific phosphodiesterase 5
(HsPDE5A; UniProt PDE5A_HUMAN: amino acids 1531-2638) was obtained from a PDE5A
cDNA clone (ORIGENE, MD, USA). Photoactivatable phosphodiesterase 4 (PhPDE4) and 5
(PhPDE5) constructs were generated by fusing the PDE4B (amino acids 630-1971) or PDE5A
(amino acids 1531-2638) with N-terminus of LAPD phytochrome (DrBPhy; UniProt
BPHY_DEIRA: amino acids 1-506). To optimize the enzymatic photoactivation, we generated a
series of variants by fusing the different length of catalytic domain of PDE4B (amino acids from
627-, 628-, 629-, 630-, 631-, 632- or 633- to 1971) or PDE5A (amino acids from 1527-, 1528-,
1529-, 1530-, 1531-, 1532- or 1533- to 2638) by site-directed mutagenesis. For protein
purification, LAPD, LAPD4 and LAPD5 were subcloned into the pCAGGS plasmid vector
(Niwa, Yamamura, & Miyazaki, 1991) with a N-terminal 6 histidine tag. For in vitro cell lysate
assay and expression in neurons, PhPDE4 and PhPDE5 were subcloned into the pCAGGS
plasmid vector with a C-terminal EGFP tag (Clontech, CA, USA). For the membrane localized
forms, a GAP43 membrane localization signal (MEM: TaKaRa Clontech, CA, USA) was
subcloned into the N-terminal PhPDE4-EGFP and PhPDE5-EGFP.
2.1.2 Protein expression and purification
HEK293 cells were transfected with plasmids using Lipofectamine 2000. The HEK293 cells
were harvested after 48 hours of transfection, and homogenized in a buffer (40 mM HEPES/Na,
pH 8.0, 0.1 mM EGTA, 5 mM magnesium acetate, 1mM DTT, and 0.01% Tween-20) by
sonication, and centrifuged at 16,000g for 15 min to clear large cell debris. The supernatant was
then isolated and used for further in vitro analysis. Using the expressed GFP, enzyme
18
concentrations were measured by GFP ELISA kit (Cell Biolabs, Inc, CA, USA). For purification,
the 6 histidine tag (6-His) constructs were expressed in HEK293 cells as above. After harvesting
and binding to Ni-nitrilotriacetic acid resin, the 6-His tag enzymes were eluted with 500 mM
imidazole according to the supplier’s instructions (Capturem His-tagged purification kit, TaKaRa
Clontech, CA, USA). The fractions that contained the enzymes were desalted and concentrated
in the above assay buffer (Amicon Ultra 30 kD, Milllipore, MA, USA). The protein
concentration of 6-His tag enzymes were determined by the Bradford assay (Bradford, 1976).
2.2 In Vitro photoactivation assay by ELISA
2.2.1 One photon illumination
In vitro photoactivation of HEK293 cell lysate containing PhPDE4 or PhPDE5 was performed
on paraffin film (Parafilm M, Sigma-Aldrich, MO, USA) covered glass slides at room
temperature in the dark. The lysate buffer contained 40 mM HEPES/Na, pH 8.0, 0.1 mM EGTA,
5 mM magnesium acetate, 1 mM DTT, 0.01% Tween-20. For each experiment, 2 µM cAMP or
cGMP was added to 100 µl of the cell lysate, then photoactivated with a 455 nm LED (5
mW/mm2; ThorLabs, NJ, USA). 60 µl of the samples were immediately added to 540 µl 0.1M
HCl to stop the reaction and the level of cAMP or cGMP was measured in triplicates for each
sample using direct ELISA biochemical assay (Enzo Life Sciences, NY, USA). All ELIZA data
was normalized to the starting amount of cAMP or cGMP at time zero and subtracted by the
empty vector control to remove any endogenous PDE activity in HEK293 cell lysate. Then the
amount of cAMP or cGMP degraded was divided by the concentration of PhPDE4 or PhPDE5 in
the lysate. Initial velocities of the enzymatic activity of PhPDEs were further divided by the
illumination time and the power of light used per surface area (mW/cm2). The duration of LED
light (0,5,15,30,60 seconds) and (0, 0.25, 0.5, 1.0, 1.5, 2.0 millisecond) was controlled by a LED
driver (DC2100, Thorlabs, NJ, USA). To examine the excitation wavelength-dependency for
LED light, we used LEDs which have various excitation peak (385, 455, 590, 660 nm, 10
mW/cm2, 30 sec, Thorlabs, UJ, USA).
19
2.2.2 Two photon illumination
For in vitro LAPD cell lysate, photoactivation of the samples were excited with focal light under
60X objective lens with two-photon laser light (30 seconds, 1,040 nm; 10 mW). For PhPDE4 and
PhPDE5 cell lysate and LAPD, PhPDE4 and PhPDE5 purified protein photoactivation, a two-
photon microscope (Custom made FV1000 MPE; Olympus, Tokyo, Japan, 60x objective lens,
NA 1.0 equipped with Spectra-Physics MaiTai HP DeepSee and InSight DeepSee) was used with
two-photon laser light (800 – 1,300 nm, 4 mW). Two-photon light dependent activity of PhPDE4
and PhPDE5 was measured at various time points (0, 2.5, 5.0, 15, 30 seconds) in the dark or after
illumination (1,100nm; 4mW). Two-photon wavelength dependency was analyzed by
illuminating lysate for 30 seconds with different wavelengths of light (800 nm, 900 nm, 1000
nm, 1,100 nm, 1,200 nm, 1,300 nm; 4 mW) (Multiphoton FVMPE-RS, 40x objective, 0.8 NA
WD 3.3mm Spectra-Physics InSight DeepSee at Scarborough imaging facility, University of
Toronto). For light intensity dependence different intensities were verified by the photodiode
power sensor (Thor Labs, S132C, Ge 700-1,800 nm, 500 mW) (5 seconds at 1, 100 nm; 0, 0.06,
0.25, 0.5, 1.0, 2.0, 4.0 mW).
2.3 Expression and Validation of PhPDEs in Neurons
2.3.1 Slice Culture
Slice culture: 1 ml of slice culture media (80 % Minimal Essential Media with 20 % horse serum
(sigma), 1 ug/ml insulin, 30 mM HEPES, 1 mM CaCl2, 2 mM MgSO4, 5 mM NaHCO3, 16.5
mM D-glucose and 0.5 mM ascorbate; pH to 7.3, 320-330 osmolarity) and one 0.4 𝜇m millicell
insert was added to each well of a 6-well plate. The plate was placed in a 37 °C, 95 % O2, 5 %
CO2 incubator. Organotypic slice cultures of rat hippocampi were prepared from postnatal day 6-
7 Sprague-Dawley rats. Rats were decapitated, following the guidelines of the university network
animal care committee approved protocol and oversight, and the whole brain removed and
placed on a petri dish containing filter paper covered with frozen, oxygenated cutting solution.
First, the brain was separated along the midline. Then the cerebellum was gently removed along
with the brainstem, midbrain and thalamus revealing the hippocampus. Using a spatula, the
hippocampus was gently scoped out and placed in ice cold dissection medium until the second
hippocampus was removed. Then hippocampi were cut into 400 𝜇m thick transverse sections.
Slices were separated in cold dissection medium and transferred onto each well of the 6-well
20
plate and incubated at 37 °C, 95 % O2, 5 % CO2. Slice culture media was refreshed every other
day (Gogolla, Galimberti, DePaola, & Caroni, 2006).
Gene gun bullet preparation: Hard tubing was dried (15 mins) using nitrogen (< 10 PSI). 100 μl
of spermidine was mixed with 12 mg of gold followed by the addition of 200 μM of DNA and
100 μL of 0.5 M cold CaCl2. Mixture was then vortexed for 6 seconds and left at room
temperature for 10 minutes. Once precipitated, tube was spun down and gold particles were
washed with 1mL of 100% ethanol 3 times. Gold particles were suspended in 1 x
polyvinylpyrrolidone (PVP). The gold-DNA-PVP mixture was drawn up with a syringe and
attached to the gene gun prep station by connecting a piece of soft tubing on the syringe to the
hard tubing on the gene gun preparation station. Gold-DNA-PVP solution was injected into the
tubing and left at room temperature for 2 minutes before the liquid was removed. Then tubing
was rotated 180° for 1 minute. Then nitrogen was turned on for 6 minutes at (0.4 pressure).
Tubing was then removed and cut into cartridges and stored in a 20 mL scintillator vial
containing Drierite (CaSO4) at -20°C (Woods & Zito, 2008).
2.3.2 Expression in Neurons
Neuron transfection: Neurons in the organotypic hippocampal slice were biolistically
cotransfected using a gene gun (120 psi helium) with DsRed and GFP-PhPDE4 or GFP-PhPDE5
or GFP-LAPD at a ratio of 1:3. Imaging and experimentation occurred 3-5 days after transfection
using an Olympus FV1000MPE based custom-made two-photon microscope (60x objective
LUMPLFLN NA 1.0, Spectra-Physics MaiTai HP DeepSee and InSight DeepSee) with 900 nm
InSight DeepSee laser excitation. Post processing of images and line scan measurements were
acquired using Metamorph software.
2.3.3 Structural Long Term Potentiation (sLTP) experiments
For sLTP experiments: slices were perfused in regular ACSF solution containing: 119 mM NaCl,
2.5 mM KCl, 4 mM CaCl2, 4 mM MgCl2, 26.2 mM NaHCO3, 1 mM NaH2PO4 and 11 mM
glucose at 30 °C equilibrated with 5 % CO2/ 95 % O2. For glutamate uncaging experiments,
ACSF was the same as above including 2.5 mM 4-methoxy-7-nitroindolinyl (MNI)-L-glutamate
with or without the addition of 50 μM forskolin.
21
sLTP was induced in the targeted dendritic spine by two photon uncaging of MNI glutamate
(720 nm laser, 4 ms duration pulses repeated at 1Hz for 60 times) the focal light point was ~1 µm
from the tip of the spine head (Kim et al., 2015). 50 µM forskolin was added into the uncaging
ACSF and applied during sLTP induction for cAMP-dependent sLTP induction (forskolin
ACSF). After uncaging the forskolin contained ACSF was returned to regular ACSF. To
photoactivate postsynaptic PhPDE4, the target spines were excited at 1,100 nm (continuous
point-scan 30 sec, 4 mW) immediately before uncaging for sLTP induction with forskolin.
Spine size change was measured by RFP (DsRed) fluorescence intensity of dendritic spines in
the three-dimensional z-stack (0.5 µm intervals) images of the dendrites typically composed of
20-30 sections using Imaris software. The average basal spine fluorescence intensity before
uncaging was defined as 100%. The spine intensity over time was subtracted from the
background intensity as well as corrected for any changes in fluorescence intensity over time of
the dendrite (arbitrary units).
2.4 Calculations and Statistical Analysis The number of replications for each experiment and statistical methods are indicated in the figure
legend. Data are presented as mean ± SEM. Statistical significance is indicated in figures as * p
< 0.05, ** p < 0.01, *** p < 0.005, and **** p < 0.001.
22
Chapter 3 Photoactivatable Phosphodiesterase 4 (PhPDE4)
Photoactivatable Phosphodiesterase 4 (PhPDE4) 3
3.1 Introduction cAMP is a critical second messenger involved in a myriad of biological processes. cAMP
is synthesized by adenylyl cyclase (AC) and degraded by a family of phosphodiesterases (PDEs).
In the brain the regulators of cAMP signaling, AC and cAMP specific PDE4 are both expressed
in regions critical for learning and memory such as the hippocampus and cortex (Kelly et al.,
2014; Xia, Choi, Wang, Blazynski, & Storm, 1993).
cAMP signaling plays an important role in synaptic plasticity that underlies learning and
memory (Brunelli et al., 1976). Learning and memory occurs through activity dependent
structural and functional changes at the synapse level known as long-term potentiation (LTP).
LTP is separated into two phase’s early-LTP (E-LTP) and late LTP (L-LTP) which coincide with
short term and long-term memory formation (Huang & Kandel, 1994; Nguyen, Abel, & Kandel,
1994). cAMP signaling during L-LTP induces critical cAMP-dependent protein synthesis
inducing long lasting functional plasticity changes. Furthermore, the application of forskolin, a
potent activator of AC, during LTP induction resulted in an enhanced enlargement of the
dendritic spine (Govindarajan, Israely, Huang, & Tonegawa, 2011). This suggests a link between
cAMP signaling and CaMKII, a major postsynaptic protein that triggers LTP and detaches from
actin enabling actin polymerization before re-associating with actin and maintaining the
enlargement of the cytoskeleton and potentiating the dendritic spine (Kim et al., 2015; Okamoto
et al., 2004). However, how cAMP is linked to this pathway inducing dendritic spine structural
changes is not clear, although cAMP signaling is important for both the structural and functional
synaptic plasticity changes, which are critical for learning and memory.
Improper regulation of cAMP signaling has been associated with numerous neurological
or neurodegenerative diseases such as Schizophrenia or Alzheimer’s disease (Gamo et al., 2013).
Specifically, a reduction in AC activity and subsequently cAMP production was found in the
brains of individuals with Alzheimer’s disease. Furthermore, the inhibition of PDE4 causing
prolonged cAMP signaling, greatly improved cognition in numerous mouse models of
23
Alzheimer’s disease (Cheng et al., 2010; Sierksma et al., 2014). Moreover, there is a common
mutation found in individuals with Schizophrenia called disrupted in schizophrenia (DISC1),
which impairs the binding of PDE4 to DISC1, which is located in dendritic spines, inhibiting
PDE4 activity in this subcellular domain (McGirr et al., 2016). This particular mutation is
associated with deficits in both working memory and long-term memory processes (Brandon,
2016; Gamo et al., 2013). This further highlights the critical role of cAMP signaling at the
synapse level and its effect on learning and memory.
Pharmacological application of forskolin or various PDE inhibitors results in widespread
alterations of cAMP signaling in the brain. This limits our understanding the role of cAMP in
LTP and learning and memory. Furthermore, these methods cannot address how cAMP alters
structural plasticity changes at a single dendritic spine level. Optogenetic approaches to produce
or suppress cAMP with light enable the precise temporal and spatial control of intracellular
signaling in living neurons.
Using our established two-photon optogenetic microscopy techniques with a modified
photoactivatable adenylyl cyclase (PAC) which produces cAMP (Stierl et al., 2011), our lab
showed that increasing post-synaptic cAMP signaling within a single dendritic spine during E-
LTP enhanced the structural plasticity and functions to regulate plasticity at nearby structures
(unpublished data). Thus, revealing a novel rapid role of cAMP signaling in structural plasticity
at the single spine level. To compliment these results and understand cAMP pathway for both
structural and functional plasticity, I will develop a novel two-photon optogenetic tool to
specifically degrade cAMP upon light illumination.
In order to generate a new optogenetic enzyme to specifically degrade cAMP, I employed
the light sensitive domain structure for LAPD but replaced the catalytic domain to the cAMP
specific PDE4 (Gasser et al., 2014). To validate the function of this optogenetic enzyme I
analyzed its photochemical properties including specificity for cAMP, light dependent activation
in vitro, wavelength dependency and light sensitivity using both one-photon and two-photon
illumination. I then applied this in living neurons for manipulating cAMP at the single dendritic
spine.
24
3.2 Results
3.2.1 Design of Photoactivatable Phosphodiesterase 4 (PhPDE4) The previously reported LAPD was developed by fusing the light sensitive phytochrome domain
from Deinococcus radiodurans to the catalytic domain of PDE2A which can bind and degrade
both cAMP and cGMP (Gasser et al., 2014). To obtain a photoactivatable PDE, which
specifically degrades only cAMP I replaced the catalytic domain of LAPD with the cAMP
specific PDE4 catalytic domain. First, I compared the catalytic domain sequences of PDE4B
with PDE2A and found the corresponding region of PDE4B where the PDE2A catalytic domain
was fused to the light sensitive bacterial phytochrome domain (PAS-GAF-PHY) for LAPD
(Gasser et al., 2014). This comparison narrowed down a potential region of the PDE4 catalytic
domain which could be used as the fusion site to join to the PAS GAF-PHY domain to create a
cAMP specific light sensitive PDE (Figure 3-1).
α-helical coiled coil structures have been commonly identified across proteins, including
PDEs, which form the linker between the regulatory domain and the catalytic domain (Pandit,
Forman, Fennell, Dillman, & Menniti, 2009). The length of this coiled coil region effects the
torsional strain between domains critical for the enzyme structure as well as activity induced
conformation changes (Hartmann, 2017). Therefore, to align the catalytic domain of PDE4 to the
coiled coil domain terminal of the phytochrome in LAPD, I chose the α-helical rich region that
can integrate with the coiled coil structure. Interestingly, the first LAPD construct (named
LAPD+2) that connected the PAS-GAF-PHY domain to the PDE2A catalytic domain had no
detectable catalytic activity upon illumination; however deletion of 2 residues created the fully
functioning LAPD (Gasser et al., 2014). Since the length of the coiled coil region connecting the
two domains is critical for optimized activity I created a series of single amino acid deletions
from the α-helical region of the PDE4 catalytic domain and fused to the PAS-GAF-PHY domain.
The fusion of this light sensitive phytochrome domain with each variant of the PDE4 catalytic
domain were named Photoactivatable PDE4 or PhPDE4. The variant number after each PhPDE4
refers to the difference of the amino acid position from the corresponding catalytic domain of
LAPD (Figure 3-2).
25
Figure 3-1. Protein sequence alignment of the C-terminal catalytic domains in PDE2A used
for LAPD and PDE4B. The fusion site for the previously reported LAPD is where the end of
the light sensitive phytochrome domain from Deinococcus radiodurans (highlighted in green)
meets the yellow highlighted region of PDE2A catalytic domain. To determine the appropriate
site of the PDE4 catalytic domain to generate a new light sensitive PDE4, I aligned the catalytic
domains of PDE4B and PDE2A. The sequence alignment provided a starting point for which
region of the PDE4B catalytic domain should be used for the new light sensitive PDE4. I also
confirmed the coiled coil domain structure using protein prediction software.
3.2.2 Optimization of the PhPDE4 light dependent photoactivation To determine if any of these fusion protein variants were successful in generating light sensitive
phosphodiesterase activity, I transfected each variant into HEK293 cells and used the cell extract
to compare the photoactivation by measuring the amount of cAMP hydrolysis using an ELISA
assay (Figure 3-2). The light stimulation induced their photo activation and PhPDE4 variant -13
demonstrated the highest light-dependent activity and was used for further characterization
experiments as PhPDE4 (Photoactivatable PDE4).
26
Figure 3-2. Design of photoactivatable PDE4 variants and their activity. (A) Depiction of the
fusion between the light sensitive phytochrome domain from Deinococcus radiodurans to the
conserved catalytic domain of PDE4 to specifically degrade cAMP. The amino acid sequences
are shown (green-segment of the phytochrome domain, yellow – portion of the PDE4 catalytic
domain). The variant numbers reflect the difference of the amino acid position from the
corresponding catalytic domain of LAPD. (B) Comparison of photoactivity between the variants
of PhPDE4. The light and dark dependent activity were measured by exposing each variant to 15
seconds in the dark or after blue light illumination (455 nm) and measuring the amount of
degraded cAMP using an ELISA biochemical assay. Data are mean ± SEM n = 9 (3 biological
replicates and 3 experimental replicates)
3.2.3 Characterization of PhPDE4 using one-photon light (LED) To examine PhPDE4 substrate specificity, I measured cAMP or cGMP dependent
photoactivation of PhPDE4. Using 455 nm blue light, PhPDE4 induced significant and rapid
degradation of cAMP but showed little effect on the level of applied cGMP, indicating rapid
activation upon illumination and cAMP specificity. The activity in the dark condition was very
limited, demonstrating light dependent activation of PhPDE4. Next I examined the response time
of PhPDE4 to light on and off. PhPDE4 photoactivation occurred in less than a millisecond
duration of light indicating rapid activation. To examine the off response of enzymatic activity, I
measured any subsequent PhPDE4 activity after the removal of light and found little additional
PhPDE4 activity after light-off, indicating rapid inactivation of PhPDE4. Along with rapid
activation, rapid deactivation upon removal of light is critical for temporal precision of this
enzyme (Figure 3-3).
27
Figure 3-3. cAMP dependent photoactivity of PhPDE4 has a rapid on and off response in
vitro. (A) Schematic of one-photon activation of PhPDE4 in vitro (B) Time course of PhPDE4
activity in vitro. The photoactivity was measured in the dark (black), or after illumination
measured by the amount of cAMP (magenta) or cGMP (blue) degraded. Data are mean ± SEM
(light+cAMP n = 4, dark+cAMP n = 3, light+cGMP n = 3). (C) Rapid time course of
photoactivation of PhPDE4. PhPDE4 was stimulated by various short millisecond durations of
light. (D) Time course of off response of PhPDE4. The off response was measured immediately
after turning off the light, after 5 seconds of illumination (plotted PhPDE4 activity to 0) and
measured any subsequent activity. Data for (C) and (D) are mean ± SEM n = 12 (4 biological
replicates, 3 experimental).
28
To examine the sensitivity of PhPDE4 to the light intensity for photoactivation, I next measured
the activity among different light intensities. The PhPDE4 activity was detected from 0.01
W/cm2 and reached a plateau by more than 0.05 W/cm2 light. The power requirement suggests
the possibility of sufficient photoactivation of PhPDE4 in the deep brain region using a LED
coupled fiber optics, which has a maximum output power of 5mW. Also, I examined the
photoactivation light wavelength-dependency of the PhPDE4. By measuring PhPDE4 activation
after exposure of various LED lights from 385 nm – 660 nm, PhPDE4 showed the broad
photoactivation of this enzyme, which was similar to the wavelength dependency with LAPD
containing the same phytochrome light sensitive domain (Figure 3-4) (Gasser et al., 2014).
According to this result, I will use 455 nm LED light for PhPDE4 photoactivation for further in
vivo studies.
Figure 3-4 PhPDE4 is activated by various light intensities and wavelengths using one-
photon light. (A) Schematic diagram representing the use of one photon blue light to measure
the intensity dependence of PhPDE4 as well as different LED to measure wavelength-
dependency of PhPDE4. (B) Light (455 nm) intensity dependence of PhPDE4 activation. The
intensity dependence of PhPDE4 was measured in vitro by exposing PhPDE4 for 15 seconds
(0.001 W/cm2 - 0.618W/cm2) (C) Wavelength dependent photo activation of PhPDE4.
Wavelength dependency was measured by activating PhPDE4 for 15 seconds in vitro using
different wavelengths of light (385, 455, 590, 660 nm; 10 mW). Data are mean ± SEM n = 9 (3
biological replicates, 3 experimental).
29
3.2.4 Two-photon characterization of PhPDE4
In order to activate PhPDE4 at the single synapse level in the brain tissue, two-photon laser
excitation is necessary as its focal volume is limited to the same size of a synapse with high NA
objective lens (~1µm3) (Denk et al., 1990). The properties of two-photon activation for LAPD,
which contains the same light sensitive phytochrome domain, have not been tested. Therefore, to
examine the two-photon wavelength dependency of PhPDE4, I measured the two-photon
excitation spectra of PhPDE4 in vitro. As the light sensitive phytochrome domain has far red
light sensitivity around 700 nm one photon excitation (Gasser et al., 2014), I used a two-photon
microscope, which covers wavelengths from 800 nm to 1,300 nm. I found the longer two-photon
excitation wavelengths 1,100-1,300nm efficiently activated PhPDE4 in vitro more than shorter
wavelengths, corresponding to the predicted two-photon excitation from the one-photon
excitation of the phytochrome domain. I used 1,100 nm two-photon excitation for further
experiments. PhPDE4 degraded cAMP in seconds using 1,100 nm illumination indicating rapid
seconds order manipulation of cAMP by two photon light. Next, to determine the necessary two-
photon excitation light intensity for the photoactivation of PhPDE4, I measured the activity in
response to various light intensities in vitro. The enzymatic activity was detected from 0.06 mW
two-photon excitation light and reached the plateau at 0.1 mW indicating sufficient two-photon
power for manipulating cAMP at synapses in the brain tissue. These results demonstrate rapid
and wavelength dependent two-photon photoactivation of PhPDE4 in vitro.
30
Figure 3-5. PhPDE4 is more efficiently activated by longer two-photon excitation and
shows rapid and light intensity dependent activation at 1,100 nm. (A) Schematic drawing of
two-photon excitation of PhPDE4 in vitro. (B) Two-photon excitation spectra for PhPDE4. 100ul
of PhPDE4 cell extract was stimulated by various wavelengths of two-photon excitation light for
30 seconds (800, 900, 1000, 1100, 1200, 1300 nm; 4 mW). (C) Time course of two-photon
photoactivation of PhPDE4 in vitro. PhPDE4 strongly degraded cAMP at 1,100 nm
(2.96±0.5μM/min/nMPhPDE4) (D) Two-photon excitation intensity dependency of PhPDE4
activation. PhPDE4 was photoactivated at 1,100 nm for 5 seconds by various two-photon
intensities (0.01 - 4.0 mW). For (B), (C), (D) data are mean ± SEM, n = 9 (3 biological
replicates, 3 experimental).
31
3.2.5 Establish soluble and membrane bound forms of PhPDE4
Endogenous PDE4 has membrane bound and soluble forms which result in important regulation
of cAMP signaling in neurons (Maurice et al., 2014). To mimic the intracellular distribution of
PDE4, I prepared a membrane bound form of PhPDE4 in addition to the soluble form by
incorporating the GAP-43 membrane localization sequence (Gauthier-Kemper et al., 2014) at the
N-terminus of the light sensitive phytochrome domain. After the photoactivation of both
membrane bound and soluble forms of PhPDE4 in vitro I found similar enzymatic activation
(455 nm, 15 seconds) of membrane bound form of PhPDE4 compared to the soluble form
(Figure 3-6). Next to observe the subcellular localization of the soluble and membrane bound
forms of PhPDE4, I biolistically co-transfected GFP-PhPDE4 with RFP (DsRed2) into
organotypic hippocampal cultured slices. The two-photon RFP fluorescence images show CA1
pyramidal neurons with dendritic spines. The GFP signal for the PhPDE4 in the dendrite was
similar to the RFP, but in the membrane bound form, GFP showed strong fluorescence intensity
at the membrane, demonstrating the expression of soluble and membrane bound PhPDE4 in CA1
hippocampal neurons (Figure 3-7).
Figure 3-6. In vitro one-photon photoactivation between membrane bound and soluble
forms of PhPDE4. (A) Schematics of domain formation of soluble and membrane bound forms
of PhPDE4. A GAP-43 membrane localization signal was inserted into the N-terminal of the
light sensitive domain. (B) Photoactivation of soluble and membrane bounds forms of PhPDE4
in vitro (455 nm, 15 seconds, 0.5 W/cm2) Data are mean ± SEM, n = 9 (3 biological replicates, 3
experimental).
32
Figure 3-7. Subcellular localization of membrane bound and soluble forms of PhPDE4 in
living neurons (A) Schematic drawing of biolistic transfection of soluble and membrane bound
forms of PhPDE4 with DsRed2. (B) Depiction of two-photon imaging of transfected CA1
pyramidal neurons in organotypic hippocampal slices for observation of subcellular localization
between soluble and membrane bound PhPDE4. (C) Representative two-photon fluorescence
live imaging of the membrane bound or soluble forms of PhPDE4. GFP (green) signal indicates
subcellular localization of PhPDE4. RFP (red) signal used as a control. (D) Soluble PhPDE4 and
(E) membrane bound PhPDE4 fluorescence profile of the white line on (C).
33
3.2.6 Validation of PhPDE4 activity in living neurons
Pharmacological application of an adenylate cyclase activator forskolin indicates that increasing
cAMP signaling enhances structural enlargement of the dendritic spine during strong synaptic
activation (Govindarajan et al., 2011) (Matsuzaki et al., 2001). Using this property, I tested
whether photo-activation of PhPDE4 can suppress the cAMP-dependent structural enhancement
of dendritic spines in living neurons. To induce cAMP dependent structural plasticity (sLTP) I
used forskolin application and two photon caged glutamate uncaging (Matsuzaki et al., 2001). 4-
Methoxy-7-nitroindolinyk-caged-L-glutamate (MNI glutamate) is a form of glutamate that is
inactive in the dark due to a photosensitive linker region blocking the glutamate function. Using
two-photon laser excitation (720nm), caged glutamate can be photolysed nearby the target spine
enabling single spine functional and structural LTP induction (10mW, 4 msec, 1 Hz, 60 times)
(Pettit D. L. et al., 1997). Using this protocol on a single targeted dendritic spines of CA1
hippocampal neurons expressing GFP-PhPDE4 and DsRed2 (volume filler), I induced structural
enlargement of the dendritic spine, called structural potentiation (sLTP) (Figure 3-8B).
Photoactivation of PhPDE4 (1, 100nm, 30sec) with the glutamate uncaging did not alter sLTP.
(Figure 3-8B)(Matsuzaki, Honkura, Ellis-Davies, & Kasai, 2004).
Next, I induced cAMP-dependent sLTP by glutamate uncaging in the presence of forskolin. The
induction of LTP with increased cAMP signaling caused further potentiated spine structure
measured at 30 minutes (Figure 3-8D). In our lab we found that postsynaptic cAMP is necessary
for structural enhancement within 30 seconds of uncaging (unpublished data). Therefore, next I
examined whether activation of PhPDE4 blocks the cAMP structural effect. To do this I photo
activated PhPDE4 using 1,100 nm for 30 seconds immediately before uncaging in the presence
of forskolin. The activation of PhPDE4 at the target spine blocked the cAMP dependent
structural enhancement, but was similar to the uncaging only sLTP (Figure 3-8D). These results
indicate that, PhPDE4 photoactivation blocked the cAMP dependent enhancement of sLTP but
not the induction of sLTP by uncaging (Figure 3-9).
34
Figure 3-8. PhPDE4 activation at single dendritic spines suppresses cAMP effect during
structural LTP (A) Depiction of sLTP induction protocol using glutamate uncaging (720 nm
excitation) and two-photon imaging (860 nm) with and without stimulation of PhPDE4. (B)
Time course imaging of baseline sLTP (black) and PhPDE4 stimulation sLTP (blue). Structural
plasticity changes at target dendritic spines were observed by measuring RFP fluorescence
intensity change over time after glutamate uncaging (720 nm, 10 mW; 4 msec, 1 Hz, 60 times)
(black) n = 11(11 spines, 9 neurons), and after PhPDE4 stimulation + glutamate uncaging (blue)
n = 10 (10 spines, 8 neurons). (C) Depiction of sLTP induction in the presence of forskolin with
and without PhPDE4 photoactivation. (D) Time course imaging of glutamate uncaging with 50
μM forskolin (magenta) n = 13 (13 spines, 8 neurons) (magenta) and the effect of PhPDE4
photoactivation in cAMP-dependent sLTP (yellow). PhPDE4 was stimulated for 30 seconds
using 1,100 nm two-photon excitation prior to glutamate uncaging with forskolin (yellow) n = 13
(13 spines, 9 neurons).
35
Figure 3-9. PhPDE4 activation at single dendritic spine causes little change during sLTP
induction but significantly suppresses the cAMP effect during cAMP dependent sLTP. (A)
Two-photon fluorescent (RFP) image of a CA1 pyramidal neuron from organotypic hippocampal
slice showing a dendrite with dendritic spines expressing GFP-PhPDE4 + RFP (DsRed2). The
while dot represents the position of glutamate uncaging at a target spine. Blue circle represents a
target spine for PhPDE4 stimulation. (B) Summary of time course imaging of sLTP without
PhPDE4 stimulation (black) and with PhPDE4 stimulation (Blue) as well as cAMP dependent
sLTP with PhPDE4 stimulation (yellow) and without (magenta). (C) Comparison of the
induction of sLTP. Structural plasticity changes between experimental groups were observed by
measuring RFP fluorescence intensity of target dendritic spines immediately after sLTP
induction (1 minute). N.S denotes not significant (P>0.05); 1 way Anova. (D) Comparison of the
potentiation of target spines. Potentiation differences between experimental groups were
observed by measuring RFP fluorescence intensity of target dendritic spines at 30minutes after
sLTP induction. **** denotes significant differences (P<0.0001) Data are mean ± SEM, n = 11
(black), n = 10 (blue) 13 (magenta), 12 (yellow).
36
3.2.7 Photoactivation of PhPDE4 for future mouse behavioral experiments
To examine whether our custom-made LED which will be implanted in the brains of freely
moving mice is sufficient to photoactivate PhPDE4, I measured the activity over various time
points. The photoactivation of PhPDE4 showed the activation in the seconds order indicating
possible suppression of cAMP signal in the brain for learning and memory experiments (Figure
3-10).
Figure 3-10. In vitro photoactivation of PhPDE4 using the implantable fiber optic LED
light source for in vivo mouse experiments (A) Schematic drawing of the implant LED in the
mouse brain and a picture of the unilateral LED light. These LEDs can also have bilateral fibers
to illuminate the PhPDE4 enzyme in both brain hemispheres. (B) In vitro time course experiment
of PhPDE4 with the custom made LED light (455 nm). Data are mean ± SEM, n = 9 (3
biological replicates, 3 experimental).
37
3.3 Summary of Chapter 3 This chapter described the development and characterization of a novel photoactivatable enzyme,
PhPDE4. This is the first optogenetic enzyme to specifically degrade the second messenger
cAMP by light. I validated the properties of photoactivation including the time-course and
substrate specificity as well as its rapid on and off response. Also, for the possibility of PhPDE4
activation in living neurons, specifically at the single dendritic spine level, I validated the
photoactivation properties with two-photon excitation light in vitro and demonstrated an
application to suppress cAMP-dependent structural plasticity (sLTP) in CA1 hippocampal
neurons. The generation of PhPDE4 will enable the investigation of cAMP signaling from the
single dendritic spine, to neural circuit and even whole brain level.
38
Chapter 4 Photoactivatable Phosphodiesterase 5 (PhPDE5)
Photoactivatable Phosphodiesterase 5 (PhPDE5) 4
4.1 Introduction The cyclic nucleotide cyclic guanosine 3’, 5’-monophosphate (cGMP) is a critical second
messenger involved in many biological processes ranging from cardiovascular, gastrointestinal
to the nervous system (Puzzo, Sapienza, Arancio, & Palmeri, 2008). cGMP is synthesized by
guanylyl cyclase (GC) and degraded by a family of phosphodiesterases (PDEs). In the brain, the
hippocampus contains a large expression of GC and cGMP specific-PDEs (Burette et al. 2002;
Szabaditis et al. 2007).
The canonical cGMP-signaling pathway, widely studied in the CA1 neurons of the
Schaffer Collateral pathway of the hippocampus during LTP induction includes NMDA receptor
mediated Ca2+ influx which stimulates Ca2+/calmodulin-activated neuronal NO synthase (nNOS)
causing NO production in the postsynaptic structure (Haley, Wilcox, & Chapman, 1992). NO
acts as a retrograde messenger and translocates into the presynaptic structure where it activates
GC to produce cGMP and initiates the increase of neurotransmitter release aiding in the
potentiation of the synapse (Hawkins, Son, & Arancio, 1998). Also, the NO activates post-
synaptic GC to produce cGMP and trigger PKG to regulate the protein synthesis, highlighting
the role of cGMP in functional plasticity and long-term memory formation (Bollen et al., 2014).
Bath application of pharmacological inhibitors of GC (LY83583), PKG (KT5823) or nNOS (3-
Br 7-Ni) blocked LTP induction in cultured rat hippocampal slices (Komsuoglu-Celikyurt et al.,
2011; Zhuo, Hu, Schultz, Kandel, & Hawkins, 1994). Furthermore, weak tetanic stimulation
does not produce LTP however the pairing of weak tetanic stimulation with injected cGMP
analogues can (Haley et al., 1992).
cGMP signaling is also involved in learning and memory processes. Application of an
nNOS inhibitor (3-Br 7-NI) or GC inhibitor (ODQ) significantly impaired both reference and
working memory (Komsuoglu-Celikyurt et al., 2011). Also, inhibition of GC and PKG impaired
memory retention in the passive avoidance task in chicks (Edwards, Rickard, & Ng, 2002).
Administration of PDE5 inhibitor, vardenafil, improved object recognition memory in different
39
memory deficit rodent models (Reneerkens et al., 2012) Furthermore, mice with PKG knockout
show deficient spatial learning using the morris water maze; a hippocampal dependent memory
process (Wincott et al., 2013). cGMP pathway has also been implicated in various brain diseases
such as Alzheimer’s disease. In Alzheimer’s disease patients, there is a strong decrease in cGMP
levels in the cerebral spinal fluid (CSF) as well as an up-regulation of PDE5 expression in the
temporal cortex (Ugarte et al., 2015) suggesting a link between cGMP signaling and disease
progression (Domek-Lopacinska, van de Waarenburg, Markerink-van Ittersum, Steinbusch, & de
Vente, 2005). Inhibition of PDE5 with sildenafil or tadalafil produced cognitive benefits in
numerous mouse models of Alzheimer’s disease (Garcia-Osta et al., 2012). Current knowledge
of cGMP signaling stems from pharmacological or genetic manipulations of cGMP-hydrolyzing
PDEs or GC activators to manipulate cGMP signaling in the brain. These approaches lack the
temporal control of local cGMP levels, which could results in off target effects or unrelated
behavioral modifications for studying the role of cGMP.
Recently we established two-photon activation of a light sensitive GC (BLGC) to
increase cGMP signaling at target dendritic spines on CA1 pyramidal neuron (Ryu, Moskvin,
Siltberg-Liberles, & Gomelsky, 2010). Utilizing this optogenetic approach, we found that an
increase in cGMP signaling suppressed the cAMP-dependent structural enhancement of dendritic
spines (sLTP) (unpublished data). This experiment revealed a novel role for cGMP signaling in
regulating structural plasticity at dendritic spines. To complement the optogenetic approach, I
developed a novel optogenetic tool to independently degrade cGMP by generating a light
sensitive PDE5. PDE5 is structurally similar to PDE2 as both contain an N-terminal tandem
GAF domain, which is similar to the bacterial phytochrome domain in LAPD (Gasser et al.,
2014; Maurice et al., 2014). The main difference between PDE5 catalytic domain and PDE2
catalytic domain is its cGMP-specific binding site (Francis, Blount, & Corbin, 2011). In order to
generate a light sensitive cGMP-specific PDE, I replaced and optimized the catalytic domain of
PDE5 into LAPD. This optogenetic tool, with BLGC, enables us to increase and decrease cGMP
signaling during LTP induction at the single dendritic spine level to understand its role during
structural and functional plasticity that underlies LTP and learning and memory.
40
4.2 Results
4.2.1 PhPDE5 enzyme design To obtain a photoactivatable PDE that specifically degrades only cGMP I replaced the catalytic
domain of LAPD (Gasser et al., 2014) with the cGMP-specific PDE5 catalytic domain. I used the
same strategy described for PhPDE4 (page 24), to determine the appropriate fusion site of the
bacterial phytochrome domain (PAS-GAF-PHY) to the catalytic domain of PDE5. To summarize
briefly, I compared the protein sequences of PDE5A with PDE2A to find the corresponding
region of PDE5 where the PDE2 catalytic domain was fused to the bacterial phytochrome
domain for LAPD (Figure 4-1). Then I used protein prediction software to determine which
residues in this region created α helices which would facilitate the formation of an α helix coiled
coil structure between the phytochrome domain and PDE5 catalytic domain (Hartmann, 2017).
Then, similar to PhPDE4 enzyme design I created a series of variants where a single amino acid
was deleted from the beginning of the PDE5 catalytic domain to optimize the length of the α-
helical region. The fusions of this light sensitive domain with each variant of the PDE5 catalytic
domain were named Photoactivatable PDE5 or PhPDE5. The variant number refers to the
difference of the amino acid position form the corresponding catalytic domain of LAPD (Figure
4-2).
41
Figure 4-1. Protein sequence alignment of the C-terminal catalytic domains in PDE2A used
for LAPD and PDE5A. The fusion site for the previously published LAPD is where the end of
the light sensitive phytochrome domain from Deinococcus radiodurans (highlighted in green)
meets the yellow highlighted region of PDE2A catalytic domain (Gasser et al., 2014). To
determine the appropriate start site of the PDE5 catalytic domain to generate a new light
sensitive PDE5 I aligned PDE5A and PDE2A. The sequence alignment provided a starting point
for which region of the PDE5A catalytic domain should be used for the new light sensitive
PDE5. Then I used protein prediction software to determine which residues of PDE5A contained
α helical structures.
4.2.2 Optimization of PhPDE5 light dependent photoactivation To determine if any of these PhPDE5 fusion protein variants were successful in generating light
sensitive phosphodiesterase activity, I transfected each variant into HEK293 cell lysate and used
the cell extract to compare the photoactivation by measuring the amount of cGMP degraded
using an ELISA assay (Figure 4-2). PhPDE5 variant -3 demonstrated the highest light-dependent
activity and was used for further characterization experiments as PhPDE5.
42
Figure 4-2. Design of photoactivatable PDE5 variants and their activity. (A) Depiction of the
fusion between the light sensitive phytochrome domain from Deinococcus radiodurans to the
conserved catalytic domain of PDE5 to specifically degrade cGMP. The amino acid sequences
are shown (green-segment of the phytochrome domain, yellow – portion of the PDE5 catalytic
domain). The variant numbers reflect the difference of the amino acid position from the
corresponding catalytic domain of LAPD. (B) Comparison of photoactivity between the variants
of PhPDE5. The light and dark dependent activity were measured by exposing each variant to 15
seconds in the dark or after one-photon illumination (455 nm) and measuring the amount of
added cGMP degraded using an ELISA biochemical assay. Data are mean ± SEM n = 9 (3
biological replicates and 3 experimental replicates)
4.2.3 Characterization of PhPDE5 using one-photon light (LED) To examine the substrate specificity of PhPDE5, I measured the amount of cGMP or cAMP
degraded after photoactivation of PhPDE5. Using 455 nm blue light, PhPDE5 induced rapid
degradation of cGMP with little effect on cAMP concentration, indicating rapid activation and
cGMP specificity. The activity in the dark condition was limited, demonstrating light dependent
activation of PhPDE5. I next examined the response time of PhPDE5 to light on and off
conditions. PhPDE5 was activated by less than a millisecond duration of light indicating rapid
activation. To examine the off response of enzymatic activity, I measured any subsequent
PhPDE5 activity after the removal of light and found little additional PhPDE5. Therefore, along
with rapid activation, I validated the rapid deactivation of PhPDE5 upon removal of light, which
is critical for the temporal precision of this enzyme (Figure 4-3).
43
Figure 4-3. cGMP dependent photoactivity of PhPDE5 has a rapid on and off response in
vitro. (A) Schematic of one-photon activation of PhPDE5 in vitro (B) Time course of PhPDE5
activity in vitro. The photoactivity was measured in the dark (black), or after illumination
measured by the amount of cAMP (magenta) or cGMP (blue) degraded. Data are mean ± SEM
(light+cAMP n = 12, dark+cAMP n = 9, light+cGMP n = 9). (C) Rapid time course of
photoactivation of PhPDE5. PhPDE5 was stimulated by various short millisecond durations of
light. (D) Time course of off response of PhPDE5. The off response was measured immediately
after turning off the light, after 5 seconds of illumination (plotted PhPDE5 activity to 0) and
measured any subsequent activity. Data for (C) and (D) are mean ± SEM n = 12 (4 biological
replicates, 3 experimental).
44
The sensitivity of PhPDE5 to different light intensities for photoactivation will provide necessary
details to determine appropriate light sources to activate PhPDE4 deep in the brains of living
animals. I measured the PhPDE5 activity in vitro in response to increasing light intensities.
PhPDE5 was activated by 0.01 mW/cm2 and reached a plateau around 0.05 mW/cm2. The low
power requirements suggest the potential to activate PhPDE5 expressed deep within brain tissue
using fiber optic LED implants. Also, I examined the wavelength-dependency for activating
PhPDE5. By measuring PhPDE5 activity after exposure to different wavelengths of LED light
(385, 455, 590, 660 nm; 10 mW) I found that PhPDE5 has broad photoactivation similar to
PhPDE4 and LAPD as they all contain the same light sensitive phytochrome domain (Figure 4-4)
(Gasser et al., 2014). Furthermore, like PhPDE4, I will use 455 nm light to activate PhPDE5 in
vivo using customized fiber optic LED implants. These results indicate that PhPDE5 is
successfully activated by one-photon light in vitro.
Figure 4-4 PhPDE5 is activated by various light intensities and wavelengths using one-
photon light. (A) Schematic diagram representing the use of one photon blue light to measure
the intensity dependence of PhPDE5 as well as different LED to measure wavelength-
dependency of PhPDE5. (B) Light (455 nm) intensity dependence of PhPDE5 activation. The
intensity dependence of PhPDE5 was measured in vitro by exposing PhPDE5 for 15 seconds
(0.001- 0.618 mW/cm2) (C) Wavelength dependent photo activation of PhPDE5. Wavelength
dependency was measured by activating PhPDE5 for 15 seconds in vitro using different
wavelengths of light (385, 455, 590, 660 nm; 10 mW). Data are mean ± SEM n = 9 (3 biological
replicates, 3 experimental).
45
4.2.4 Two-photon characterization of PhPDE5
Two-photon laser excitation has a focal volume around ~1µm3 with high NA objective lens
which can localize activation of PhPDE5 at a single targeted dendritic spine in deep brain tissue
(Denk et al., 1990). Therefore, similar to PhPDE4, I determined the two-photon wavelength
dependency of PhPDE5 to determine best excitation spectra for photoactivation. As expected,
PhPDE5 two-photon excitation spectra closely resembles the PhPDE4 excitation spectra as they
both contain the same light sensitive phytochrome domain (Gasser et al., 2014). Specifically,
PhPDE5 was strongly activated by longer two photon wavelengths of light (1,100 nm -1,300 nm)
in vitro. I used 1,100nm excitation for further experiments and found that PhPDE5 degraded
cGMP in seconds, indicating the ability to manipulate by two-photon light. Next to determine the
necessary two-photon excitation light intensity for photoactivation of PhPDE5, I measured the
activity in response to various light intensities in vitro. The enzymatic activity was detected from
0.06mW two-photon excitation light and reached the plateau at 0.1 mW indicating sufficient
two-photon power for manipulating cGMP at synapses in the brain tissue. These results indicate
that PhPDE5 is successfully activated by two-photon light in vitro.
46
Figure 4-5. PhPDE5 is more efficiently activated by longer two-photon excitation and
shows rapid and light intensity dependent activation at 1,100 nm. (A) Schematic drawing of
two-photon excitation of PhPDE5 in vitro. (B) Two-photon excitation spectra for PhPDE5. 100
ul of PhPDE5 cell extract was stimulated by various wavelengths of two-photon excitation light
for 30 seconds (800, 900, 1000, 1100, 1200, 1300 nm; 4 mW). (C) Time course of two-photon
photoactivation of PhPDE5 in vitro. PhPDE5 strongly degraded cGMP at 1,100 nm (D) Two-
photon excitation intensity dependency of PhPDE5 activation. PhPDE5 was photo activated at
1,100 nm for 5 seconds by various two-photon intensities (0.01 mW - 4 mW). For (B), (C), (D)
data are mean ± SEM, n = 9 (3 biological replicates, 3 experimental).
47
4.2.5 Establish soluble and membrane bound forms of PhPDE5
cGMP signaling is regulated by both soluble and membrane bound PDEs which results in
important regulation of cGMP signaling in neurons (Menniti, Faraci, & Schmidt, 2006). Using
the same strategy described for PhPDE4 (page 40) I generated a membrane bound version of
PhPDE5 using the same GAP-43 membrane localization sequence. I found similar enzymatic
activation in vitro (455 nm, 15 seconds) of membrane bound form of PhPDE5 compared to the
soluble form (Figure 4-6). Next to observe the subcellular localization of the soluble and
membrane bound forms of PhPDE5, I biolistically co-transfected GFP-PhPDE5 with RFP
(DsRed2) into organotypic hippocampal cultured slices. The two-photon RFP fluorescence
images show CA1 pyramidal neurons with dendritic spines. The GFP signal for PhPDE5 in the
dendrite was similar to the volume filler RFP, but for the membrane bound form, GFP showed
strong fluorescence intensity just at the membrane, demonstrating the expression of soluble and
membrane bound PhPDE5 in CA1 hippocampal neurons (Figure 4-7).
Figure 4-6. In vitro one-photon photoactivation between membrane bound and soluble
forms of PhPDE5. (A) Schematics of domain formation of soluble and membrane bound forms
of PhPDE5. A GAP-43 membrane localization signal was inserted into the N-terminal of the
light sensitive domain. (B) Photoactivation of soluble and membrane bounds forms of PhPDE5
in vitro (455 nm, 15 seconds). Data are mean ± SEM, n = 9 (3 biological replicates, 3
experimental).
48
Figure 4-7. Subcellular localization of membrane bound and soluble forms of PhPDE5 in
living neurons (A) Schematic drawing of biolistic gene gun transfection of soluble and
membrane bound forms of PhPDE5 with DsRed2. (B) Depiction of two-photon imaging of CA1
pyramidal neurons in organotypic hippocampal slices expressing PhPDE5 for observation of
subcellular localization between soluble and membrane bound forms. (C) Representative two-
photon fluorescence live imaging of the membrane bound or soluble forms of PhPDE5. GFP
(green) signal indicates subcellular localization of PhPDE5. RFP (red) signal used as a control.
(D) Soluble PhPDE5 and (E) membrane bound PhPDE5 fluorescence profile of the white line on
(C).
49
4.2.6 Photoactivation of PhPDE5 for future mouse behavioral experiments
To examine whether PhPDE5 can be sufficiently activated in the brains of freely moving mice
using our custom-made implantable LEDs, I first measured the activity of PhPDE5 in vitro over
various time points using this light source. The photoactivation of PhPDE5 occurred in the
seconds order indicating possible suppression of cGMP signal in the brain for future learning and
memory experiments (Figure 4-8).
Figure 4-8. In vitro photoactivation of PhPDE5 using the implantable fiber optic LED light
source for in vivo mouse experiments (A) Schematic drawing of the implant LED in the mouse
brain and a picture of the unilateral LED light. These LEDs can also have bilateral fibers to
illuminate the PhPDE5 enzyme in both brain hemispheres. (B) In vitro time course experiment of
PhPDE5 with the custom made LED light (455 nm). Data are mean ± SEM, n = 9 (3 biological
replicates, 3 experimental).
4.3 Summary of Chapter 4
This chapter paralleled Chapter 3, describing the development and characterization of a novel
photoactivatable enzyme PhPDE5. This is the first optogenetic enzyme to specifically degrade
the second messenger cGMP. Similar to PhPDE4, I validated the properties of PhPDE5
photoactivation including the time course and substrate specificity and its rapid on and off
response. I also validated the two-photon photoactivation excitation properties in vitro and
examined the diffuse and membrane bound expression in vivo. The generation of PhPDE5 now
enables us to suppress cGMP signaling at a targeted dendritic spine as well as independently
block cGMP signaling in living animals during learning and memory tasks.
50
Chapter 5 Comparison of photo activity between PhPDEs and LAPD
Comparison of photo activity between PhPDEs and 5LAPD
5.1 Introduction
All three light sensitive enzymes including a previously reported LAPD (Gasser et al.,
2014) to degrade cAMP and/or cGMP are composed of the light sensitive phytochrome domain
from Deinococcus radiodurans fused to the catalytic domain of PDE2, PDE4 or PDE5 (Gasser et
al., 2014). Due to the substrate specificity of the catalytic domain, this strategy was successful
for engineering three different light sensitive PDEs which all retain their cAMP/cGMP
specificity. There are 11 families of PDEs, which are classified according to their substrate
specificity, amino acid sequences, and regulatory processes (Bender & Beavo, 2006). PDEs are
modular enzymes containing an N-terminal regulatory domain and a C-terminal catalytic domain
to hydrolyze cAMP or cGMP or both.
This chapter will compare the in vitro light sensitive activity of LAPD, PhPDE4 and
PhPDE5 with regular PDE activity. Also, I further characterized LAPD using two-photon
excitation to verify in vitro activation as well as the diffuse expression of this enzyme in vivo.
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5.2 Results
5.2.1 Comparison of the light sensitive activation between LAPD, PhPDE4 and PhPDE5.
To compare the previously reported light sensitive activity of LAPD (Gasser et al., 2014),
alongside PhPDE4 and PhPDE5, using our in vitro techniques, I illuminated cell lysate
containing LAPD using blue light (455 nm), which caused rapid degradation of both cAMP and
cGMP (Figure 5-1). The light dependent cAMP and cGMP hydrolysis are relatively lower than
PhPDE4 and PhPDE5.
Figure 5-1. In vitro photoactivation of PhPDE4, PhPDE5 and LAPD on cAMP and cGMP
levels. Summary of optogenetic toolkit, which degrades cAMP and/or cGMP using 455 nm light.
(B) Comparison of the initial activity in vitro of PhPDE4, PhPDE5 and LAPD with cAMP
(magenta) or cGMP (blue) plotted per min per W/cm2. The initial velocity of PDE activity is
plotted per minute per W/cm2. PhPDE4 activity for cAMP vs cGMP (p = 0.0004 ****), PhPDE5
activity for cAMP and cGMP (p = 0.0066**), and LAPD activity cAMP vs cGMP (p = 0.3, ns).
Statistical analysis: Mann Whitney test, one tailed.
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5.2.2 Two-photon activation of LAPD
In order to activate LAPD specifically at the single synapse level of living neurons in the brain
tissue, two-photon laser illumination is necessary. Due to the different properties of single
photon and two-photon light, I validated LAPD activation using two-photon illumination. I
found that LAPD degraded cAMP and cGMP after 15 seconds of 1,040 nm illumination,
indicating strong photoactivation by two-photon laser light (Figure 5-2).
Figure 5-2. Two-photon photoactivation of LAPD efficiently induces the degradation of
both cAMP and cGMP. (A) Schematic drawing of two-photon excitation of LAPD in vitro. (B)
Comparison of LAPD activity for cAMP and cGMP in vitro. The photoactivity of LAPD with
1,040 nm two-photon light. Data are mean ± SEM, n = 9 (3 biological replicates, 3
experimental).
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5.2.3 Photoactivation properties of purified LAPD, PhPDE4 and PhPDE5
To determine if the kinetics of PhPDE4 and PhPDE5 optogenetic enzymes are similar to the
previously reported LAPD under one-photon (660 nm) and two-photon excitation (1, 100 nm), I
measured their photochemical properties using purified proteins and compared the
photoactivation efficiency among these optogenetic tools. PhPDE5 has slightly higher
photoactivation compared to PhPDE4 and LAPD using both one photon and two photon
illumination (Table 1).
Table 1. Photoactivation of purified LAPD, PhPDE4 and PhPDE5 using one-photon and two-photon excitation light.
5.2.4 Establish soluble and membrane bound forms of LAPD
PDE2 has membrane bound and soluble forms which results in important regulation of cAMP
and cGMP signaling in neurons (Bender & Beavo, 2006). Therefore, along with soluble LAPD,
to prepare a membrane bound form of LAPD, I examined the addition of two different
membrane localization sequences to the N terminal of the phytochrome domain: the PDE2
membrane localization sequence (MLS) and the GAP-43 membrane localization sequence. I
biolistically co-transfected GFP-LAPD with RFP (DsRed2) into organotypic hippocampal
cultured slices. The GFP signal for the soluble LAPD in the dendrite showed similar distribution
to the RFP, but for the membrane bound forms, GFP showed strong fluorescence intensity at the
membrane for GAP-43 and aggregated expression for MLS membrane bound form (Figure 5-3).
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Figure 5-3. Subcellular localization of membrane bound and soluble forms of LAPD in
living neurons (A) Schematic drawing of gene gun transfection of soluble and membrane bound
forms of LAPD with DsRed2. (B) Depiction of two-photon imaging of CA1 pyramidal neurons
in organotypic hippocampal slices expressing LAPD for observation of subcellular localization
between soluble and membrane bound forms. (C) Representative two-photon fluorescence live
imaging of the LAPD-mem, LAPD-MLS or LAPD-soluble. GFP (green) signal indicates
subcellular localization of LAPD. RFP (red) signal used as a control. (D) Soluble LAPD and (E)
LAPD-mem fluorescence profile of the white line on (C).
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5.2.5 Preparation of LAPD lenti viral construct for in vivo applications
In order to use this optogenetic enzyme as well as PhPDE4 or PhPDE5 in vivo to study the effect
of degrading cAMP and/or cGMP level, I used lenti virus system with CaMKII promoter, which
restricts expression to the excitatory neurons (Wang et al., 2013; Feng Zhang et al., 2010). I
expressed and validated the CaMKII-GFP-LAPD plasmid in HEK293 cells by analyzing the
expression of GFP using confocal microscopy. Next I confirmed the photoactivity of this
construct in HEK293 cells (Figure 5-4).
Figure 5-4. Validate expression and light dependent activity of pLenti-LAPD in HEK293
cells. (A) Lentivirus under the CaMKII promoter will be used to limit expression of LAPD to
excitatory neurons in the hippocampus. (B) HEK293 cells were transfected with CaMKII-GFP-
LAPD plasmid. Fluorescent images verified transfection efficiency. Images were obtained using
a confocal microscope (10X objective, 475 nm excitation). Scale bar represents 50 μm. (C)
Photoactivity of CaMKII-GFP-LAPD. Using 455 nm light, LAPD photoactivation was
confirmed in the presence of 4 μM of cAMP after 2 minutes of illumination (p<0.05, n = 3). Data
(n = 3). Statistical significance was calculated using paired t-test.
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5.3 Summary of Chapter 5 This chapter described the in vitro activation of LAPD to compare with PhPDE4 and PhPDE5. I
confirmed the one-photon photoactivation of LAPD as well as the two-photon photoactivation.
To compare their photochemical properties, I purified LAPD, PhPDE4 and PhPDE5 and
measured their photoactivation kinetics under one-photon and two-photon light stimulation in
vitro. I also expressed the soluble form of LAPD and the membrane bound form of LAPD in
CA1 hippocampal neurons and validated their localization in vivo. Furthermore, for future mouse
behavioral experiments I validated a lentiviral construct in vitro to restrict LAPD expression to
hippocampal neurons. The generation of PhPDE4 and PhPDE5 along with LAPD will enables us
to study cAMP/cGMP signaling at a targeted dendritic spine as well as deep in the brain tissue of
living animals during learning and memory tasks.
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Chapter 6 Discussion
This chapter is divided into 6 sections. Section 6.1 is a summary of the project. Section 6.2
discusses photoactivation properties of PhPDEs. Section 6.3 discusses in vivo application of
PhPDE4 in living neurons. 6.4 Highlights the comparisons of kinetics and limitations of these
new optogenetic tools. Section 6.5 discusses future work and important neuroscience questions
that can be addressed using these tools to conclude the thesis.
Discussion 6
6.1 Summary of the project We established the optogenetic toolkit for manipulating cAMP and cGMP with the necessary
spatiotemporal precision to study their roles in neuronal activity underlying learning and
memory. This cyclic nucleotide optogenetic toolkit contains the previously reported bacterial
photoactivatable adenylyl cyclase (bPAC) which produces cAMP in response to blue light (Stierl
et al., 2011), the blue light sensitive guanylyl cyclase which produces cGMP (Ryu et al., 2010)
and a far red light sensitive, light activatable phosphodiesterase (LAPD) which degrades both
cAMP and cGMP (Gasser et al., 2014).
In this thesis, I established two optogenetic enzymes, PhPDE4 and PhPDE5 that degrade second
messenger molecules, cAMP and cGMP, independently upon light illumination. These
optogenetic tools provide us with great spatial and temporal control of cAMP and cGMP
signaling in an unprecedented way. By controlling the light-dependent activation, this enables us
to study endogenous cAMP/cGMP functions from millisecond to minutes order to understand
their rapid role in synaptic functions and related learning and memory. Furthermore, I optimized
two-photon laser light-dependent activation of PhPDE4 and PhPDE5 in vitro and validated the
function at single synapse level in living neurons. Thus, PhPDE4 and PhPDE5 will provide novel
powerful tools to study cAMP/cGMP signaling from the synapse level to the brain’s cognitive
functions.
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6.2 Photoactive properties of PhPDEs By fusing the light sensitive domain from Deinococcus radiodurans to the catalytic domain of
PDE4 or PDE5 I generated two new light sensitive enzymes named photoactivatable PDE4
(PhPDE4) and photoactivatable PDE5 (PhPDE5) that independently degrade cAMP and cGMP
respectively. This basic design stemmed from LAPD where illumination caused the degradation
of both cAMP and cGMP (Gasser et al., 2014). Utilizing the similarity of a regulatory domain of
PDEs (GAF-B) and a light sensitive phytochrome domain (PHY), the phytochrome domain was
tightly connected to the catalytic domain of PDEs by linking their helical structures (Gasser et
al., 2014). Therefore, the predicted light-dependent conformational changes for photoactivation
are similar to the activation of PDEs (Figure 6-1).
Figure 6-1. Schematic diagram of the predicted photo cycle of PhPDEs. (A) Closed state
when no light is present. (B) Illumination induces conformational changes enabling (C) cAMP or
cGMP to bind to the catalytic domain causing their degradation. (D) Removal of light then
rapidly reverts the enzyme back into its closed conformational state impairing further
degradation of cAMP or cGMP.
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Under dark conditions, the conformation of the enzyme renders the binding site for cAMP or
cGMP on the catalytic domain blocked (Figure 6-1;A). Once illuminated, the light-sensitive
phytochrome domain absorbs photons of light inducing a rotation of the chromophore resulting
in a conformational change and catalytic activity (Figure 6-1;B). It is believed that this rotation
causes rearrangement of hydrogen bonds enabling the evolutionarily conserved PHY domain to
convert from B-sheet confirmation into an alpha helix (Ziegler & Moglich, 2015). This change in
phytochrome domain causes conformational changes in the attached catalytic domain resulting in
the opening of the enzyme and presenting the binding site for cAMP or cGMP (Figure 6-1; C).
As the catalytic domain of phosphodiesterases are highly conserved each contains a glutamine
binding pocket specific for either cAMP or cGMP, I hypothesized that the substrate specificity
should be maintained without the presence of their regulatory domain (Zhu, Yang, Dai, & Huang,
2013). In in vitro photoactivation experiments, I detected that PhPDE4 rapidly degrades cAMP
but not cGMP, and PhPDE5 rapidly degrades cGMP but not cAMP (Figure 6-1;C) (see figure 3-
3, 4-3). I also validated that PhPDE activity can be induced broadly by various one photon light
sources (385-660 nm) and is sensitive to low light intensities (0.01-0.05mW/cm2) (see Figure 3-
4, 4-4). Moreover this activation can occur within 0.25 milliseconds of illumination suggesting
the application for studying rapid cAMP/cGMP functions (see figure 3-3, 4-3).
PhPDEs are also sensitive to two-photon laser light illumination. This is critical to
degrade cAMP and/or cGMP at a single dendritic spine to study their role in synaptic plasticity
as two-photon focal points form within a micron excitation volume, similar to the size of a
dendritic spine, when using high numerical aperture (NA) objective lenses, making this
technique suitable for manipulating light-sensitive enzymes at the single synapse level deep
within living brain tissue (Denk et al., 1990). I found that the longer two-photon excitation
wavelength, which corresponds to the single photon excitation wavelength efficiently, activated
PhPDE4, PhPDE5 and LAPD (see figures 3-5, 4-5, 5-2). I also confirmed that after the removal
of light, catalytic activity of these light sensitive enzymes ceased (Figure 6-1;D) (see figure 3-3,
4-3). Thus, PhPDE4 and PhPDE5 serve as the first optogenetic enzymes to independently
degrade cAMP or cGMP by light.
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6.3 In vivo validation of PhPDEs in living neurons To verify that these tools can be applied for neuroscience research, at first I expressed PhPDE4
and PhPDE5 as well as LAPD in living neurons (see figures 3-7, 4-7 and 5-3). All three light
sensitive PDEs showed homogenous expression in CA1 pyramidal neurons of hippocampal
slices. As PDEs have membrane bound forms in addition to soluble forms, I prepared new
membrane bound forms of light sensitive PDEs. Endogenous PDE2A localizes to synaptosomal
membranes, therefore to generate membrane bound forms I first inserted the membrane
localization sequence from this PDE to LAPD. However, this construct caused aggregated
expression throughout the hippocampal neurons. I next tried a commonly used membrane
localization sequence from Growth Associated Protein 43 (GAP-43), which attaches to the
neuron membrane by dual palmitoylation targeting GAP-43 to lipid rafts (Gauthier-Kemper et
al., 2014). Using GAP-43 membrane localization sequence, LAPD was successfully localized to
the membrane and the same strategy was applied for PhPDE4 and PhPDE5 (see figures 3-7, 4-7
and 5-3).
To validate the functionality of these optogenetic enzymes in vivo, I applied PhPDE4 to validate
the role of cAMP signaling on structural plasticity. During LTP induction, there is an increase in
the size of the dendritic spine, which persists over 30 mins called structural LTP (sLTP). We
found that post-synaptic cAMP enhances sLTP. By reducing the cAMP signaling by
photoactivation of PhPDE4 at the target spine during cAMP-dependent sLTP, PhPDE4
successfully blocked this cAMP-dependent enhancement (See figure 3-8, 3-9). Since the
difference between PhPDE4, PhPDE5 and LAPD is the catalytic domain containing the substrate
specific site for cyclic nucleotide-binding, these results suggest the functions of PhPDEs in living
systems.
6.4 Comparisons and limitations of PhPDE4, 5 and LAPD
To compare the optogenetic enzymes to the previously published LAPD, I measured
photoactivation of LAPD, PhPDE4 and PhPDE5 under one-photon and two-photon illumination
using the purified proteins (see table 1). Comparing the reported Vmax of endogenous PDE2,
PDE4 and PDE5 activity to the activity of LAPD, PhPDE4 and PhPDE5 some major differences
61
arise. Strikingly, the activity of LAPD was dramatically lower for both cAMP and cGMP
compared to the PDE2 activity of 123 µM cGMP/min/mg and 120 µM cAMP/min/mg,
suggesting that photoactivation of LAPD does not fully activate the PDE2 catalytic domain
(Rosman et al., 1997). While, PhPDE4 activity was slightly higher than the known PDE4 range
(0.0078-0.4 µM/min/mg) (Huston et al., 1997), and PhPDE5 activity was only slightly lower
than the known PDE5 range (1.0-1.3 µM/min/mg) (Loughney et al., 1998). This similarity
suggests that photoactivation of PhPDEs produces an almost maximally activated enzymatic
state. Moreover, it has been reported that endogenous PDE5 has higher enzymatic activity than
PDE4, which is a trend that I saw with PhPDE4 and PhPDE5. Also, PDE2 has higher activity for
cGMP degradation than cAMP degradation that is also retained in the light sensitive activity of
LAPD. Therefore, the optogenetic enzymes, despite being engineered to only turn on upon
illumination still retain similar enzymatic properties compared to endogenous form. These results
suggest that the photo activation of PhPDE4 and PhPDE5 in living neurons has the potential to
mimic the same level of activity as endogenous PDEs.
One limitation to PhPDEs is their broad spectrum of activation by one-photon illumination.
Although the phytochrome domain from Deinococcus radiodurans is known as a far red light
sensitive domain, I found that PhPDE4 and PhPDE5 were sensitive to a broad range of one-
photon wavelengths (see figures 3-4, 4-4) (Gasser et al., 2014). For example during
electrophysiology experiments on hippocampal slices, one could co-transfect PhPDE4 and PAC
(blue light sensitive adenylyl cyclase) with the intent to increase and decrease cAMP signaling in
the same cells using different wavelengths of light. Using this example, while PhPDE4 could be
independently activated using 660 nm light without stimulating PAC, but activation of PAC
using 455 nm blue light will also activate PhPDE4. In order to achieve this type of
experimentation, optogenetic enzymes with narrow wavelength specificity and far enough
spectral separation will be required. In contrast, spectral differences using two-photon
illumination such that PhPDE4 and 5 were best activated by more than 1,100 nm two-photon
wavelengths, whereas PAC and BLGC are activated by 700-800 nm two-photon excitation. This
could be a possibility for independent activation of PhPDE4 with PAC for regulation of cAMP
and PhPDE5 with BLGC for cGMP using two-photon illumination. Co-expressing these pairs
would enable the activation or suppression of cAMP or cGMP signaling in a single dendritic
spine during synaptic plasticity.
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6.5 Future Applications It is well established that both cAMP and cGMP are important for long term memory through the
activation of CREB and generation of plasticity related proteins. PhPDE4 or PhPDE5 will enable
us to suppress endogenous cAMP/cGMP functions to study the roles of cAMP/cGMP in
cognitive function. To validate the functionality and correct optogenetic setup to activate
PhPDE4 and PhPDE5 in living animals for future behavioral experiments, the novel object
recognition test will be used to observe the effect of blocking cAMP or cGMP on memory
formation. After this validation, PhPDE4 or PhPDE5 can be stimulated with varying spatial and
temporal dimensions during novel object recognition for short or long term memory, or 5 choice
serial reaction time task for attentional memory to understand the effect of suppressing the
spatiotemporal cAMP or cGMP signaling on cognitive output. Furthermore, these enzymes can
be used to suppress the rapid cAMP or cGMP functions during LTP or LTD induction using
electrophysiology on rodent hippocampal slices. Finally, using our established two-photon
optogenetic approach, I will monitor the effect of suppressing cAMP or cGMP signaling at
single targeted dendritic spines on structural plasticity.
6.6 Conclusion cAMP and cGMP are universal second messengers for intracellular signaling critical for a
myriad of biological functions. cAMP and cGMP have been implicated in synaptic plasticity and
their misregulation has been associated with a variety of neurodegenerative disorders.
Pharmacological, genetic and electrophysiology techniques have shown the correlative role of
cAMP and cGMP in learning and memory. To further examine the role of cAMP or cGMP
during this activity dependent process requires a method to manipulate their signaling within a
biologically relevant timescale while restricting such activation spatially as to leave neighboring
cells unaffected. Optogenetic tools enable perturbations to each of the three levels of neural
activity with this spatiotemporal accuracy. To complement previously characterized PAC and
BLGC, which produce cAMP and cGMP respectively, I have created and validated two new
optogenetic tools to independently suppress cAMP or cGMP signaling for the first time using
light. The completion of this optogenetic toolkit will enable the accurate dissection of cAMP and
cGMP during activity dependent synaptic plasticity.
63
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