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Developing novel in vitro model systems to investigate therapeutic hypotheses for peroxisome biogenesis disorders

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Developing novel in vitro model systems to investigate therapeutic hypotheses for peroxisome biogenesis disorders

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Content DEVELOPING NOVEL IN VITRO MODEL SYSTEMS TO INVESTIGATE
THERAPEUTIC HYPOTHESES FOR PEROXISOME BIOGENESIS DISORDERS
by
Bradford H. Steele
A Thesis Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
MASTER OF SCIENCE
(BIOCHEMISTRY AND MOLECULAR BIOLOGY)
May 2019
Copyright 2019 Bradford H. Steele

ACKNOWLEDGMENTS

I would first like to express my deepest gratitude to my mentor, Dr. Joseph Hacia, for his
support throughout my Master’s project and his dedication to the field of peroxisome biogenesis
disorders. His commitment to helping patients and families affected by this disease is truly
remarkable and inspirational. I also would like to thank the other members of my committee, Dr.
Michael Stallcup and Dr. Baruch Frenkel. Their advice and support has proven invaluable during
this process and I am grateful to have had such a knowledgeable, kind and supportive committee.
I would also like to thank all the members, past and present, of the Hacia lab for their
support and aid in my research endeavors. Namely, I would like to express my special thanks to
Leone D’Antonio for his hard work and dedication to the project and his friendship through my
time in the Hacia lab.
Lastly, I wish to express my gratitude to my family and friends, who have always been
my foundation and my inspiration to reach farther than I thought possible for myself. Their love,
support and belief in me is an irreplaceable gift.

TABLE OF CONTENTS
CHAPTER 1: INTRODUCTION ................................................................................................... 1
CHAPTER 2: LITERATURE REVIEW ........................................................................................... 8
Metabolic functions ............................................................................................................................. 8
Peroxisome assembly ........................................................................................................................ 12
CHAPTER 3: VALIDATION OF PEX1-MUTANT HEPG2 CELLS FOR PEROXISOME BIOGENESIS
DISORDERS ............................................................................................................................ 15
Abstract ......................................................................................................................................... 15
Introduction ................................................................................................................................... 16
Materials and methods .................................................................................................................. 20
Cell lines and cell culture ................................................................................................................... 20
Confirmatory analysis of mutations by DNA sequencing .................................................................. 21
Immunofluorescence ......................................................................................................................... 22
Results ........................................................................................................................................... 23
Identification of PEX1-null cell lines .................................................................................................. 23
Immunoflourescence assessment and chemical treatment response .............................................. 23
Discussion ...................................................................................................................................... 24
CHAPTER 4: USING CRISPR-CAS9 BASE-EDITOR TO CREATE AND CORRECT PEX1-p.G843D
MUTATION ............................................................................................................................ 26
Abstract ......................................................................................................................................... 26
Introduction ................................................................................................................................... 27
Materials and Methods .................................................................................................................. 32
gRNA Cloning ..................................................................................................................................... 32
Cell Culture ........................................................................................................................................ 33
Plasmid Transfection ......................................................................................................................... 33
Immunofluorescence ......................................................................................................................... 33
Cell isolation ...................................................................................................................................... 34
PCR and Sequencing .......................................................................................................................... 34
Subcloning ......................................................................................................................................... 35
Results ........................................................................................................................................... 35
Generation Of The PEX1 c.2528G>A p.G843D Mutation in HEK293T Cells ....................................... 35
Correction of PEX1c.2528G>A p.G843D in patient immortalized primary fibroblasts ...................... 36
CHAPTER 5: FUTURE DIRECTIONS .......................................................................................... 40
HepG2 PEX1-null CRISPR-modified cell lines ................................................................................... 40
BE3 PEX1 Editor (PEX1-p.G843D mutation generation) ................................................................... 41
ABE Pex1 Editor (PEX1- p.G843D mutation correction) ................................................................... 42
In vivo base editing ........................................................................................................................ 42
REFERENCES .......................................................................................................................... 45

1
CHAPTER 1: INTRODUCTION

Peroxisomes are organelles present in virtually all cells of eukaryotic organisms that play
vital roles in numerous metabolic pathways [1-3]. Thus, their proper assembly and function are
essential to human health and development. Depending on their metabolic needs, a human cell can
contain anywhere from a few hundred to several thousand peroxisomes. Peroxisomes are single-
membrane bound, usually spherical in shape and range in size from 0.1 to 1 micron in diameter
[4]. Nevertheless, peroxisomes in different cell types have been noted to vary in their
morphological appearance from round to rod-shaped to tubular and network-like [5, 6].
Peroxisomes are one of the more recently discovered organelles in eukaryotic cell biology
[7]. They were first formally recognized and characterized by Nobel-prize winning cytologist and
biochemist, Christian de Duve, in 1966 [8]. The de Duve research group isolated peroxisomes
from rat liver tissue and discovered that several H
2
O
2
-producing oxidases and H
2
O
2
-degrading
enzymes co-localized in the matrix of this new organelle, which he later dubbed the “peroxisome”
[8].
The peroxisome is characterized by a proteinaceous matrix surrounded by a single
membrane studded in transport proteins [9]. The matrix proteins perform enzymatic tasks essential
for particular metabolic pathways, many of which are not performed anywhere else in the cell [10,
11]. Namely, these tasks include beta-oxidation of very long chain fatty acids ((VCLFA, 24-28
carbon atoms in length), branched chain fatty acids (including phytanic and pristanic acid), and
pipecolic acid [10, 11]. They also play critical roles in the biosynthesis of cholesterol, bile acids,
docosahexaenoic acid (DHA) and ether-phospholipids, including plasmalogens [10, 12]. Based on
the physical manifestations of patients with peroxisomal disorders, peroxisomes are essential for
the normal structure and functions of the endocrine, digestive, nervous, renal, reproductive,

2
respiratory, and skeletal systems [13]. Other important roles of the peroxisome include the
detoxification of glycolate to glycine and the deactivation of certain reactive oxygen species
(ROS), including hydrogen peroxide [10] and reactive nitrogen species (RNS) [14].
Peroxisomes recruit and house a unique set of membrane and enzymatic proteins that are
essential to their proper function. In humans, there are an estimated 70 unique proteins that will
ultimately end up in the peroxisome membrane or matrix [1] (Table 1). Proteins destined for the
location in the peroxisomal membrane or matrix are initially translated in the cytoplasm and bear
a Peroxisomal Targeting Sequence (PTS) [15]. There are two unique signal types, PTS1, which is
always found on the C-terminal end of the protein and is the most common signal amongst
peroxisomal proteins, and the PTS2 signal [16], which lies near the N-terminus in the protein
sequence. The PTS1 and PTS2 signals are recognized by two separate peroxisomal proteins, PEX5
and PEX7 respectively [17]. These two proteins play key roles in recognizing and recruiting
cytoplasmic peroxisome-bound proteins to the membrane of the peroxisome [17].
The importance of proper peroxisome function is highlighted by a number of multi-
systemic disorders in humans which result from inherited mutations in peroxisome-related genes
[1, 13, 18]. Peroxisomal disorders can be broadly categorized into two major groups: 1)
peroxisome biogenesis disorders (PBDs), which affect the complete assembly of the fully
functional peroxisome and 2) mutations in enzymatic peroxisome proteins or particular transport
proteins essential to peroxisome function [19]. In the first, the inability of the cell to assemble fully
functional peroxisomes leads to a dramatic halt in several metabolic pathways whose steps are
normally carried out in peroxisomes. In the second category, an enzyme essential to a particular
pathway is mutated and therefore cannot carry out its metabolic role, while the peroxisome as a
whole and most other pathways are still kept intact.

3
Table 1. Peroxisomal matrix proteins and their targeting sequences (from ref [10])
Peroxisomal protein Gene Symbol Signal Sequence
Peroxisomal beta-oxidation
Acyl-CoA oxidase 1 ACOX1 PTS1 -SKL
Acyl-CoA oxidase 2 ACOX2 PTS1 -SKL
Acyl-CoA oxidase 3 ACOX3 PTS1 -SKL
L-bifunctional protein EHHADH PTS1 -SKL
D-bifunctional protein HSD17B4 PTS1 -AKL
Peroxisomal beta-ketothiolase 1 ACAA1 PTS2 -RLQVVLGHL
Peroxisomal beta-ketothiolase 2 SCP2 PTS1 -AKL
Alpha-methylacyl-CoA racemase AMACR PTS1 -(K)ASL
Carnitine acetyltransferase CRAT PTS1 -AKL
Carnitine octanoyltransferase CROT PTS1 -THL
Delta 3,5-, delta 2,4-dienoyl-CoA isomerase ECHI PTS1 -SKL
Peroxisomal 2,4-dienoyl-CoA reductase 2 DECR2 PTS1 -AKL
Peroxisomal 3,2-trans-enoyl-CoA isomerase PEC1 PTS1 -SKL
Very-long-chain acyl-CoA synthetase SLC27A2 PTS1 -LKL
Acyl-CoA thioesterase 2 PTE1 PTS1 -SKL
Acyl-CoA thioesterase 1B PTE2 PTS1 -SKV
Peroxisomal trans-2-enoyl-CoA reductase PECR PTS1 -AKL
Peroxisomal alpha-oxidation
Phytanoyl-CoA 2-hydroxylase PHYH PTS2 -RLQIVLGHL
2-Hydroxyphytanoyl-CoA lyase HPCL2 PTS1 -(R)SNM
Plasmalogen biosynthesis
Dihydroxyacetone phosphate acyltransferase GNPAT PTS1 -AKL
Alkyldihydroxyacetone phosphate synthase AGPS PTS2 -RLRVLSGHL
Fatty acyl-CoA reductase 1 MLSTD2 — —
Fatty acyl-CoA reductase 2 MLSTD1 — —
Glyoxylate metabolism
Alanine:glyoxylate aminotransferase AGXT PTS1 -KKL
Lysine metabolism
L-pipecolate oxidase PIPOX PTS1 -AHL
Oxygen metabolism
Catalase CAT PTS1 -(K)ANL
Peroxiredoxin V (PMP20) PRDX5 PTS1 -SQL
D-amino acid oxidase DAO PTS1 -SHL
D-aspartate oxidase DDO PTS1 -(K)SNL
Glycolate oxidase HAO1 PTS1 -SKI
Hydroxyacid oxidase 2/3 HAO2/3 PTS1 -SRL
Epoxide hydrolase EPHX2 PTS1 -SKM
Glutathione S-transferase class Kappa GSTK1 PTS1 -ARL
Polyamine metabolism
N1-acetylspermine/spermidine oxidase PAOX PTS1 -(R)PRL

4

While X-linked adrenoleukodystrophy (X-ALD), caused by mutations in the ABCD1 gene
that encodes a lipid transporter in the peroxisome outer membrane, is the most common
peroxisomal disorder [20-22], PBDs collectively account for a majority of the remaining
peroxisome related disorders seen in patients. The PBDs themselves can be divided into Zellweger
spectrum disorder (ZSD), the focus of this thesis, and rhizomelic chondrodysplasia punctata
(RCDP) [1, 13, 18]. Of these, ZSD is the more prevalent and occurs at a rate of about 1 in 50,000
births in North America. Classically, prior to knowledge of their molecular underpinnings, ZSDs
were categorized into three specific groups based on their severity. The most severe form of the
disease was referred to as Zellweger syndrome (ZS), wherein patients rarely survive beyond the
first year after birth [1, 13, 18]. Patients with this form of the disease have severe congenital brain
malformations caused by impaired neuronal migration and development, as well as
hypomyelination. The disease can also affect the function of several other organ systems, leading
to hepatomegaly (enlarged liver), chondrodysplasia punctata (punctate calcification of cartilage in
various regions of the body), vision and hearing loss, and renal cysts. About half of ZSD patients
fall into the other two classical categories of the disease, neonatal adrenoleukodystrophy (NALD)
and infantile Refsum disease (IRD) [1, 13, 18]. These milder forms of disease lead to progressive
vision and hearing loss, mild to moderate intellectual disabilities, liver dysfunctions and other
medical conditions, with patients often surviving into early adulthood (Table 2). This family of
diseases is characterized by increases and decreases in several metabolic byproducts which can be
used for diagnosis, which eventually is supplemented with cell culture characterization and genetic
testing (Table 3).

5
Table 2. Clinical features of ZSD by severity and age of onset (reference [23])
Features Neonate 1-6 months 6 months – 4 years >4 years
Neuronal migration defects S
Chondrodysplasia punctata S
Renal cortical cysts S
Respiratory compromise S
Craniofacial dysmorphism S, I, M I, M I, M
Jaundice S, I, M I
Hepatomegaly, liver dysfunction
cholestasis
S I, M I, M I, M
Hypotonia, failure to thrive S,I I, M I, M M
Sensorineural hearing loss S I, M I, M M
Cataracts S I, M I, M M
Retinal degeneration S I, M I, M M
Psychomotor retardation S I, M I, M M
Seizures S I I M
Leukodystrophy I I M
Adrenal insufficiency I, M M
Osteopenia I I, M
Calcium oxalate renal stones I, M I, M
Peripheral neuropathy M M
Cerebellar ataxia, atrophy M M
Amelogenesis imperfecta I,M I, M
Abbreviations for clinical severity classes: S, severe; I, intermediate; M, mild

Table 3. Current diagnostic tests for PBDs (reference [23])
Test Metabolite/gene PBD
Plasma VLCFA ↑C26:0, C26:1 & C24/C22, C26/C22 ratio ZS, NALD, IRD
Plasma phytanic and pristanic acid Increased
ZS, NALD, IRD,
RCDP1 (phytanic only)
RBC plasmalogens Reduced
ZS, NALD, IRD,
RCDP1
Urine/plasma bile acid intermediates ↑di- and tri-hydroxycholestanoic acid ZS, NALD, IRD
Urine/plasma pipecolic acid Increased ZS, NALD, IRD
Cultured fibroblasts Confirm metabolites by enzyme assays
ZS, NALD, IRD,
RCDP1
Molecular genetic testing
PEX 1- 3, 5, 6, 10, 11b, 12- 14, 16, 19,26
PEX7
ZS, NALD, IRD
RCDP1
Abbreviations: ZS, Zellweger Syndrome; NALD, Neonatal Adrenoleukodystrophy; IRD, Infantile Refsum Disease;
RCDP1, Rhizomelic Chondrodysplasia Punctata Type 1

6
To date, treatments for ZSDs are limited and palliative in nature, with the clinical value of
FDA-approved bile acid replacement therapies still under discussion [24-30]. Nevertheless, initial
work by the laboratory of Dr. Nancy Braverman (McGill University) [31] and then in collaboration
with the Hacia laboratory have identified candidate small molecules that improve peroxisome
assembly in human cell model systems carrying the most common ZSD mutation, PEX1-p.G843D,
a hypomorphic allele that encodes a PEX1 protein with residual activity, as opposed to a null
mutation which produces a protein with no residual activity or a protein that is not expressed [32].
Small molecules have been identified via cell-based high-content screening (HCS) of chemical
libraries, which has proven to be an effective means of discovering potential therapeutic candidate
drugs. The screening system developed by Dr. Braverman and employed in the Hacia laboratory
is based on monitoring the localization of a modified green fluorescent protein with a C-terminal
PTS1 sequence (GFP-PTS1) into peroxisomes of ZSD patient-derived immortalized patient
fibroblasts. In untreated ZSD patient cells, the GFP-PTS1 reporter protein is expected to be
cytoplasmic due to compromised peroxisome assembly. Nevertheless, in the presence of small
molecules that rescue peroxisome assembly, the GFP-PTS1 reporter protein becomes punctate. As
such, the rescue of peroxisome assembly via small molecule treatments can be observed via
fluorescence microscopy. Although quite powerful, this assay system is limited in its ability to
identify the rescue of downstream peroxisome functions and also only represents one of numerous
other cell types. The overall goal of the Hacia laboratory is to development more effective targeted
therapies for ZSD. As one major approach we are undertaking to realize this long-term goal, we
are actively seeking to develop new cell-based assays which can better detect rescue of peroxisome
function in a high-throughput manner. As part of this research initiative, we seek to develop and
optimize new cell lines for screening that are more representative of different tissue types most

7
affected by disease in patients. This will also enable us to begin to explore the potential
effectiveness of candidate small molecules in various organ systems and furthermore provide
novel insights into the cell-type specificity of peroxisomal functions.

8
CHAPTER 2: LITERATURE REVIEW

As noted in Chapter 1, peroxisomes are membrane-bound organelles that play pivotal roles
in several anabolic and catabolic pathways, as well as in the disposal of harmful byproducts of
metabolism. They are essential to human health and the normal function of multiple organ systems.
In mammalian cells, there are an estimated over 80 peroxisomal proteins that are vital to
peroxisomal assembly and division, import of lipid substrates into the peroxisome, and also the
enzymatic activities that are performed within the peroxisome matrix. As previously mentioned,
the loss or reduction of function of the peroxisome-associated proteins can leads to severe
developmental and degenerative disorders collectively referred to as PBDs. Below, we will begin
to provide a broad overview of what is currently known about peroxisome assembly, structure,
and functions.
Metabolic functions

Peroxisomal Fatty Acid Alpha-Oxidation

While most fatty acid metabolism, such as the beta-oxidation of long chain, medium, and
short chain fatty acids, takes place in the mitochondria, some types of fatty acid can only be
metabolized in the matrix of the peroxisome. This includes the branched-chain fatty acids
(BCFAs), which usually contain a methyl group at the C3-position. BCFAs make up only a small
percentage of ingested fatty acids from the diet. The most prominent BCFA is phytanic acid which
is obtained through the consumption of dairy products, ruminant animal fats and from certain fish
[33-35]. BCFAs have also been suggested to play an important role in establishing the microbiota
of the newborn gastrointestinal tract [36]. These fatty acids cannot undergo catabolism via beta-
oxidation due to the presence of a functional group on their beta-carbon. The process of alpha-

9
oxidation of BCFAs involves the linkage of a CoA to the terminal carboxyl group, the
hydroxylation of the alpha-carbon and finally the removal of the terminal carboxyl-CoA group.
This process produces a substrate of length n-1, moving the branched carbon out of the beta-
position so that beta oxidation may then occur.
Peroxisomal Fatty Acid Beta-Oxidation

Other types of fatty acids that cannot be metabolized in the mitochondria are the very long
chain fatty acids (VLCFAs), waxy fatty acids containing 22 or more carbons. These fatty acids
undergo a special type of beta-oxidation that only occurs in the peroxisome and involves the
production of hydrogen peroxide as a byproduct. Other lipids, such as pristanic acid (a byproduct
of phytanic acid alpha-oxidation) are also metabolized in the peroxisome. Additionally, the
peroxisome beta-oxidation system is vital for the production of some polyunsaturated fatty acids.
ROS metabolism

A number of peroxisomal oxidases necessary for metabolic processes are present in the
matrix of the peroxisome. These enzymes produce several reactive oxygen species (ROS) as
byproducts, such as hydrogen peroxide. The highly reactive nature of ROS poses a danger to the
cell and so must be deactivated quickly within the peroxisome. Therefore, the peroxisome also
contains a number of enzymes, such as catalase, glutathione peroxidase and peroxiredoxin V
(PMP20), which decompose hydrogen peroxide and other ROS [3]. The ROS produced by
peroxisomes have also been linked to modulating the activity of signaling pathways through ATM
and the MTORC1 [37-39].

10

Plasmalogen Biosynthesis

Plasmalogens are a type of ether-phospholipid that are found in numerous organs and
tissues, with particular enrichment in the nervous, immune and cardiovascular systems [40].
Although their functions are not fully defined, plasmalogens have been suggested to play roles in
protecting the cell from the damaging effects of ROS and may act as signaling molecules and
modulators of membrane dynamics. They are an important component of the myelin sheath,
making up about 70% of the ethanolamine glycerophospholipids found in myelin [41]. As
exemplified by the peroxisomal disorder rhizomelic chondrodysplasia punctata, plasmalogen
deficiencies can have widespread effects on the health and development of the nervous system
[42-44].
Plasmalogen biosynthesis involves the activity of multiple organelles, but is initiated in the
peroxisome [10]. The process involves the interaction of two peroxisomal
enzymes, GNPAT (glycerone phosphate acyl transferase) and AGPS (alkyl-glycerone phosphate
synthase) on the luminal side of the peroxisome membrane [10]. The process begins with
dihydroxyacetone phosphate (DHAP), which is acylated at the sn-1 position by GNPAT.
Subsequently, the acyl group is exchanged for an alkyl group by AGPS. This byproduct is further
modified in the peroxisome and the endoplasmic reticulum (Figure 1). The inability of the cell to
assemble fully functional peroxisomes, where GNPAT and AGPS can associate and function
together, has drastic effects on plasmalogen biosynthesis. This in turn leads to defects in nervous
tissue function and development, as well as complications in other organ systems.

11

Figure 1. Plasmalogen structure and biosynthesis. (A) Chemical structure of plasmalogens.
Plasmalogens are structurally similar to diacylglycerophospholipids but differ at the sn-1 position where
the acyl group has been replaced with a vinyl ether bond in plasmalogens. (B) Plasmalogen biosynthesis.
The biosynthesis of plasmalogens begins in the cytosol with the conversion of a fatty acid to a fatty alcohol
(step 1). Within the peroxisome, DHAP is acylated by GNAP (step 2). This product then has its acyl group
replaced with an alkyl group and is covalently bonded to a fatty alcohol (step 3). All further plasmalogens
modification occur in the endoplasmic reticulum (step 4). From reference [1].

12
Glyoxylate Metabolism
Peroxisomes also play a role in glyoxylate metabolism [10]. The peroxisomal enzyme
alanine:glyoxylate aminotransferase (AGT), found primarily in liver peroxisomes, converts
glyoxylate to glycine to prevent the build-up of oxylate, an end product of glyoxylate
decomposition. The loss of function of AGT or inability of the cell to form peroxisomes can lead
to the build-up of oxylate crystals in the kidney and liver, leading to failure of these organ systems
[45-47].
Peroxisome assembly

The classic model of peroxisome assembly involves two pathways: 1) peroxisomes are
generated de novo via vesicular budding off of the ER, after which they undergo a maturation
process in which peroxisomal enzymes are imported into the matrix and 2) new peroxisomes are
formed from mature peroxisomes via fission. More recently, evidence has shown that vesicles
originating from the mitochondria are also involved in the de novo pathway, fusing with the
vesicles budding off of the endoplasmic reticulum. These mitochondria-derived vesicles contain
matrix proteins and peroxisome membrane proteins (PMPs) which aid in the maturation process
(Figure 2).

13

Figure 2. Diagram of peroxisome de novo and fission formation pathways. The de novo
pathway of peroxisome formation involves the budding of vesicles from both the endoplasmic
reticulum and mitochondria, which later fuse to form pre-peroxisomes. These vesicles already
contain some transport membrane proteins essential for peroxisome maturation. The maturation
process involves the transport and accumulation of peroxisome matrix and membrane proteins.
Mature peroxisomes can then form new daughter peroxisomes via fission. Figure taken from
reference [48].

The transport of proteins destined for the peroxisome is highly selective and mediated by
the presence of peroxisome-specific amino acid sequences on peroxisomal matrix proteins. There
are two known peroxisome targeting signal (PTSs). The majority of proteins destined for the
peroxisome matrix bear an extreme C-terminal tripeptide signal (PTS1) with consensus sequence
(S/A/C)-(K/H/R)-(L/M) [15] . The PTS1 signal is recognized and bound by PEX5, which then
mediates the transport of the protein into the matrix through interactions with a membrane-bound
transport system. The second signaling motif (PTS2), found only on three peroxisome proteins
(AGPS, ACAA1, and PHYH), is a less conserved, N-terminal internally located octapeptide signal
with consensus sequence (R/K)-(L/V/I)-(X)5-(H/Q)-(L/A) [16]. This signal is recognized by

14
PEX7, which can later bind PEX5 once it has bound its cargo, ultimately leading to a similar
transport process as with proteins bearing PTS1.

15
CHAPTER 3: VALIDATION OF PEX1-MUTANT HEPG2
CELLS FOR PEROXISOME BIOGENESIS DISORDERS

Abstract

Peroxisome biogenesis disorders affect the body globally, although some organ systems
are more seriously affected. The majority of patients with ZSD suffer from liver disease. This is
due to the fact that hepatocytes, the primary cell type found in the liver, play significant roles in
several metabolic pathways, including glycogen storage, lipid and serum protein biosynthesis,
metabolism of many dietary lipids, and detoxification of xenobiotics [49]. The high levels of lipid
metabolism occurring in hepatocytes makes their peroxisome an especially vital organelle and, as
such, hepatocytes generally contain a much higher number of peroxisomes than other cell types.
Currently, due in part to the fact that they are readily accessible and easy to maintain, the most
common cell-based models for ZSD are derived from patient and genetically engineered mouse
fibroblasts. Although these cells have proven invaluable for screening small molecule libraries, we
would prefer to be working with cell-based models that more closely represent affected cell-types.
Obtaining primary hepatocytes from patients is not feasible for a number of important ethical and
technical considerations. We hypothesized that the widely utilized HepG2 liver-cancer
immortalized cell-line carrying common ZSD mutations could serve as a better model for testing
the effects that drugs we have identified may have on liver function and health and better elucidate
the mechanisms by which the rescue is occurring. To do this, we employed CRISPR/Cas9 gene
editing to introduce PEX1 mutations in HepG2 cells.

16

Introduction

ZSD is a disease continuum with phenotypes ranging from mild to severe, depending on
the gene mutated and the effect of the that mutation on the function of the downstream protein.
Although varying in severity, most ZSD patients have liver disease, variable neural-developmental
delay, retinopathy and perceptive deafness at birth [13, 50]. Since ZSD is a relatively simple
Mendelian disorder, the genetics of this spectrum of disease has been well elucidated via genetic
sequencing technology; however, the exact effect that lack of peroxisome assembly and function
have on cell and tissue development are not well understood. Moreover, because of the differences
in the prominence of peroxisomes, organelle crosstalk, and differential gene expression between
varying cell types, the mechanisms of disease may vary from one cell type to another [51]. As
previously mentioned, our current cell-based model for screening small molecule libraries and
monitoring peroxisome assembly involve cells derived from ZSD patient fibroblasts. Although
useful for initial screens and identification of potential small molecules, we feel it would be of
special valuable to work with cell models that more closely represent the organ systems most
effected by disease. Because the liver is a highly effected organ [24-29, 52], we endeavored to
establish a cell model using the widely used HepG2 liver cancer cell line and introduce genetic
mutations to these cells via CRISPR-Cas9.
Hepatocytes are the main functional and structural cell type of the liver, making up about
70-85% of the mass of the liver as a whole. These cells are responsible for several vital metabolic
processes, including glycogen storage, lipid, bile salt, and protein biosynthesis, dietary lipid
metabolism and detoxification of drugs or other toxic substances introduced to the body. For these
reasons, the study of hepatocyte function and health is considered the gold standard for
understanding the mechanisms of liver disease. Be this as it may, obtaining sufficient numbers of

17
primary ZSD hepatocytes for experimental purposes is not possible due to ethical concerns, limited
availability, complicated isolation procedures and short life span of cells. Thus, no immortalized
hepatocyte cell line has ever been available as a model for ZSD liver disease. To overcome this
shortcoming, we considered the possibility of using already established and immortalized liver-
derived cell lines as an ideal alternative model to study liver diseases pathophysiology and to
perform drug screens.
HepG2 are an immortalized hepatocellular carcinoma cell line originally derived from liver
tissue from a 15-year old male [53]. HepG2 cells have been utilized in numerous studies for
purposes including the effects of hypoxic conditions on the liver [54], cytochrome p450 induction
by xenobiotics [55], predicting drug-induced liver injury [56], investigating the anti-cancer effects
of small molecules [57], and for screening the hepatotoxicity of newly discovered, potentially
therapeutic chemicals at early stages of drug development [58]. The ability to generate specific
genetic modifications to the genome via CRISPR-Cas9 presents a promising opportunity to
establish novel in vitro cell cultures models for PBD-ZSD that better reflect the cell type specificity
of disease. HepG2 cells carrying mutations relevant to PBD-ZSDs could be a powerful and
significant tool for further elucidating the role of peroxisome assembly and function in this disease
spectrum and may help in drug discovery efforts.
ZSD arise through the disruption of peroxisome biogenesis as a result of defective
peroxisomal proteins essential to import into either the peroxisome membrane or its matrix.
Defects in these proteins are caused by mutations in the peroxin (PEX) genes. Of these genes, the
most commonly mutated gene found in ZSD patients is PEX1, which encodes an AAA (ATPase
associated with diverse cellular activities) – ATPase, and accounts for the genetic defect in about
two-thirds of PBD-ZSD patients [59]. The PEX1 protein interacts with the PEX6 and PEX26

18
protein to form a membrane-anchored exportomer complex that recycles PEX5 back into the
cytoplasm using ATP hydrolysis as an energy source (Figure 3) [60-63]. When this complex
cannot function properly, the PEX5 protein remains embedded in the peroxisome membrane and
cannot return to the cytoplasm to perform its duty of binding PTS1-bearing cytoplasmic
peroxisome proteins destined for the peroxisome [17, 64, 65]. This results in the inability to import
vital proteins into the peroxisome, thus making the peroxisome nonfunctional.

Figure 3. Overview of peroxisome matrix protein import. (Step 1) PTS receptor binding.
Peroxisome matrix protein import involves both PEX5 and PEX7 which bind their ligands, the
PTS1 and PTS2 signals, respectively. PEX7 requires binding to a larger isoform of PEX5 (PEX5L)
to import its cargo. (Step 2) Docking. The cargo loaded complex binds PEX13 and PEX14 which
are embedded in the peroxisome membrane. (Step 3) Matrix enzyme translocation. Cargo is
unloaded into the peroxisome matrix via formation of pore created by PEX5 and PEX14. (Step 4)
Receptor recycling. The membrane-embedded PEX5 is monoubiquitinated by the PEX2/10/12
complex. The monoubiquitinated PEX5 is then removed from the membrane by the exportomer
complex (PEX1/6/26), driven by ATP hydrolysis. The ubiquitin is subsequently removed from
PEX5 to allow a new cycle of import to begin. PEX7 is returned to the cytosol in an ATP-
independent manner. Note that PEX1 function is essential for returning PEX5 to the cytosol for
proper import to occur. Figure taken from reference [23].

19
Currently, there are well over 40 known mutations that have been reported in PBD-ZSD
patient cohort studies, representing a wide spectrum of ancestral backgrounds [66, 67]. Of these
mutations, about 20-30% of patients carry at least one copy of the PEX1 c.2528G>A p.G843D
allele, a hypomorphic mutation that results in a misfolded PEX1 with reduced function associated
with milder clinical phenotypes [66, 67]. As compared to wild-type PEX1, the interaction between
PEX1-p.G843D and PEX6 is reduced to less than 70%, resulting in impaired peroxisomal import
[66, 67].
The clustered regularly interspaced short palindromic repeat (CRISPR)-CRISPR-
associated Cas9 system for genetic modification in living cells has proven to be an extremely
powerful tool in the field of biology since its characterization and introduction as a laboratory tool
in 2013 [68-70]. This system allows for the precise and efficient editing of virtually any locus in
the genome and has been widely used to inactivate, modify and insert genetic information into the
genome of several organisms, including human cells. The modern CRISPR-Cas9 technology is an
engineered version of a bacterial adaptive immune system, the type II CRISPR-Cas system. Type
II CRISPR-Cas allows bacteria to protect themselves from invading phage and plasmid DNA by
incorporating small pieces of exogenous DNA into a specific CRISPR locus in their genome.
These small fragments are then transcribed, resulting in small RNA (crRNA) fragments
complimentary to the invading DNA, which hybridize with a larger RNA stem-loop structure
(tracrRNA). These hybrid crRNA:tracrRNA then complex with a Cas protein, which acts as an
endonuclease. When the same foreign DNA is present, the RNA-Cas complex can recognize and
cleave the foreign DNA, thereby protecting the bacteria. This simple system has been engineered
to be used in virtually any organism to specifically target and cleave genomic DNA. The
crRNA:tracrRNA has been engineered into a single RNA sequence and a guide RNA (gRNA) of

20
choice can be cloned into it using traditional plasmid cloning strategies. This system has provided
an inexpensive, efficient and simple means of genetically modifying living cells and organisms,
dramatically transforming biologic research in recent years. Some estimates place CRISPR-Cas9
mediated genome editing at an efficiency of 80% on the targeted locus, which is as high or higher
than other means of editing such as TALENS. One concern with CRISPR-Cas9 is the possibility
of off-target editing, since the system can tolerate mismatches in gRNA-genomic DNA
complementarity. This is a notable problem and must be monitored when using CRISPR-Cas9 to
edit the genome. Much research is being done to address this issue and reduce the amount of off-
target editing.
The traditional CRISPR-Cas9 technology can efficiently introduce double-strand breaks
(DSBs) in genomic DNA at a predetermined locus in the genome. The applications of this are
many. Generation of DSBs can be used to produce gene knock-out or mutants by formation of
indels via the error-prone nonhomologous end-joining (NHEJ) pathway, knock-in of specific
mutations or entire genes via the homology-directed repair (HDR) pathway, or to produce tumor-
associated chromosomal translocations.
We therefore hypothesized that the traditional CRISPR-Cas9 genome editing system could
be used to generate HepG2 cells bearing PEX1 null alleles to serve as better in vitro models for
future PBD-ZSD studies. Herein, we describe the generation and characterization of CRISPR-
Cas9 edited HepG2 PEX1 cell lines.
Materials and methods

Cell lines and cell culture
HepG2 cells were acquired from the Coriell Institute Cell Repositories (CIRC) and cultured
at 37°C with 5% CO
2
with Fibroblast medium (high-glucose DMEM supplemented with 10% FBS,

21
penicillin/streptomycin, vitamin solution, essential and nonessential amino acids, all from Thermo
Fisher Scientific).
To generate mutants, GenScript (Piscataway, NJ) was contracted to carry out the CRISPR-
Cas9 mediated editing of our HepG2 cells to produce PEX1 mutants. First, cells were transfected
with the CRISPR-Cas9 system using a gRNA targeting an exonic region of the PEX1 locus. After
validating from bulk transfected cells that editing had occurred, we received the cells from
Genscript. These cells were then plated on 96-well plates through a series of ten-fold serial
dilutions, with a final calculated seed density of 1 cell/well. Isolated cells were expanded and later
analyzed by Sanger sequencing for presence of PEX1 mutations.
Confirmatory analysis of mutations by DNA sequencing

Because the product obtained from Genscript consisted of a mixed population of HepG2
cells possibly containing cells bearing either no PEX1 mutation or various deletion mutants of
PEX1, it was necessary to isolate and expand single cells to generate a homogenous population of
cells with the same mutation in order to maintain consistency for downstream experiments.
Because of their relevance to disease, we were most interested in producing cells homozygous for
PEX1 knockout mutations. Genomic DNA was extracted from isolated populations after clonal
expansion. Primers flanking the targeted locus located within exon 15 of PEX1 were used to PCR
amplify the region for use in Sanger sequencing. The following primers were used for this
purpose:
gPEX1e14-1-Forward: 5’- CACTATAGATTTGTCAACCTGATTTTC -3’
gPEX1e14-2-Forward: 5’- CAGAGTATATCCACCAGAGA -3’
gPEX1e15-1-Reverse: 5’- TTGAGACCTCACTCTGTCAT -3’
gPEX1e15-4-Reverse: 5’- CAACAAGTGTTTACTGAGTTACCA -3’

22
PCRs were performed as 20uL reactions containing 150 ng of gDNA, 5uL of Phusion HF
5X Buffer (Thermo Fischer Scientific), 0.5 uM final concentration of each primer, 0.2 mM final
dNTP concentration, and 0.4 units of Phusion polymerase (Thermo Fischer Scientific). An initial
denaturation at 98°C was performed, followed by thirty five cycles of 98°C for 10 seconds, 60-
65°C for 30 seconds, and 72°C for 1-3 minutes (depending on expected size of product) and a final
five minute extension at 72°C. No DNA controls were run alongside these to monitor for DNA
contamination. PCR products were analyzed on 1% agarose gels to confirm the presence of
appropriately sized fragments. These products were then subcloned into the pCR
®
Blunt II-TOPO
®

vector (Thermo Fisher Scientific) and subsequently transformed by heat-shock into chemically
competent E. coli cells (Thermo Fisher Scientific). Individual colonies were picked and expanded
in LB broth overnight. Plasmid DNA was extracted from these expanded colonies via PureLink
Miniprep (Thermo Fisher Scientific) which was sent for sequencing to detect presence of
mutations on each allele.
Immunofluorescence

To visualize peroxisome assembly, PEX1-mutant HepG2 cells were transduced with a
lentivirus expressing a GFP-PTS1 which will be imported into healthy peroxisomes. HEK293 cells
were kindly provided by Dr. Wange Lu’s laboratory for lentivirus packaging. To produce lentivirus
in HEK293 cells, 5 µg for each of the following were transfected: pMD2.G (expressing VSV-G
envelope protein), psPAX2 (expressing Pol and Gag) and pCT-Pero-GFP (lentiviral vector
designed to express GFP-PTS1 reporter gene, from System Biosciences). Vectors were co-
delivered into HEK293 cells using Lipofectamine
®
LTX (Thermo Fisher Scientific) in 100-mm
dishes. After 36 hours post-transfection, lentivirus-containing medium was harvested filtered
through a 0.45 µm cell media filter. Lenti-X™ Concentrator (Clontech) was then used to

23
concentrate the lentivirus which was then resuspended in fibroblast medium. HepG2 cells were
transduced twice with lentivirus expressing GFP-PTS1 reporter [71].
Results
Identification of PEX1-null cell lines

Twenty cell lines were successfully isolated and expanded. Of these, seven were subjected
to subcloning and sequencing. Cell line number 1 was identified as PEX1 c.2522delA/2519del14,
meaning it was compound heterozygous where one allele had a single base deletion while the other
had a 14-bp deletion in the targeted exonic region. As such, both alleles represent frameshift
mutations. Another cell line, number 7, was found to be homzoygous for the PEX1 c.2522delA
frameshift mutation. The other colonies were either heterozygous or contained undesired
mutations. Further characterization was performed on these cell lines number 1 and 7.

Immunoflourescence assessment and chemical treatment response

These two cell lines were then transduced with our GFP-PTS1 lentivirus, along with
unaltered HepG2 cells. As expected, in unaltered HepG2 cells, many robust puncta, representing
heathly peroxisomes with functional import, could be seen in all cells with little cytoplasmic GFP.
Contrary to unaltered HepG2, both of our CRISPR modified cells showed very little to no punctate
GFP-PTS1 reporter and instead all GFP-PTS1 reporter was cytoplasmic, indicating a lack of
functioning peroxisomes (Figure 4).

24

Figure 4. HepG2 and PEX1-mutant HepG2 cells expressing GFP-PTS1. Unedited HepG2 cells
and CRISPR-edited PEX1-null cell lines expressing GFP-PTS1 reporter protein (green) and
counter-stained with DAPI (blue). HepG2 cells show robust punctate GFP-PTS1 signal while both
mutant cell lines show diffuse, cytosolic GFP-PTS1 expression, suggesting an impaired ability to
correctly assemble peroxisomes.

Discussion

All forms of disease caused by peroxisome dysfunction are associated with liver disease,
highlighting the importance of peroxisomes for normal liver function. HepG2 cells serve as a
promising cell culture model system for liver disease due to their similarities to hepatocytes, albeit
with some limitations. Here, we generated two HepG2 cell lines which are homozygous or
compound heterozygous for null PEX1 mutations. Sequencing data shows no presence of wild
type PEX1, suggesting we have obtained pure, isolated cell lines from our bulk CRISPR modified
HepG2 cells. Additionally, via immunoflourescence, we observe no functional peroxisome import
using our GFP-PTS1 reporter gene as refernece. This suggests that these cells do not express a
viable PEX1 protein and therefore cannot form fully functional peroxisomes due to impaired
import. More characterization is necessary to fully validate these cells, including Western blot,
Real-time PCR analysis of PEX1 mRNA expression, and biochemical lipid profiling via liquid
tandem mass spectometry (LC-MS/MS).

25
Overall, these cells will allow us to further investigate the pathomechanisms of liver
disease in PBDs. They may also be valuable in future drug screening studies, allowing for
validation of candidate small molecules in a liver-model system. These cells have already been
shared with colleagues working in the field of peroxisome biology who have confirmed in their
own labs that our HepG2 cells do not have peroxisomal import and are beginning to use the cells
for their own research purposes.

26
CHAPTER 4: USING CRISPR-CAS9 BASE-EDITOR TO
CREATE AND CORRECT PEX1-p.G843D MUTATION

Abstract

Since its introduction as a biological research tool in 2013 [68-70], CRISPR-Cas9 genome
editing technology has revolutionized both basic and translational biological science [72, 73]. In
this short time, researchers have tinkered with the Cas9 protein to create systems that are capable
of performing a multitude of previously impossible tasks. In addition to its wild-type function of
acting as an endonuclease, researchers have been able to engineer various forms of the Cas9 protein
via directed evolution that do not create double-strand breaks but instead produce single-strand
nicks (Cas0-nickase) or do not cut the DNA strand at all (dead-Cas9). Furthermore, researchers
have experimented with creating Cas9 fusion proteins to bring specific protein functionality to a
targeted region of the genome. Examples of these include dead-Cas9 fused to GFP to map and
visualize the location of genomic loci and fusions to DNA activators and repressors to enhance or
decrease expression of a targeted gene respectively.
Recently, several research labs, including David Liu’s at Harvard, have begun producing
Cas9 proteins fused to various DNA/RNA base editors [74, 75]. These Cas9-base editors are
capable of converting bases within a window of the targeted protospacer to produce specific
mutations. These Cas9-base editors function by producing a single-strand nick in the targeted
region, promoting the recruitment of DNA repair machinery, and simultaneously bringing the base
editor in proximity of the cut site to induce a base conversion and repair process. This technology
holds great promise in its ability to precisely modify the genome without introducing double-strand
breaks, which can form insertions or deletions (indels) at the targeted locus and any other off-
target regions with similar protospacer sequence. Here, we have implemented these systems to 1)

27
create the PEX1 c.2528G>A p.G843D mutation in cells for the purpose of creating new cell models
and 2) repairing the PEX1 c.2528G>A p.G843D mutation in cells that already carry it to explore
the system’s efficiency and efficacy for possible clinical usage in the future.

Introduction
The most common mutation found in ZSD patients is the PEX1 c.2528G>A p.G843D
allele, representing about 30% of cases [66, 67]. This is a hypomorphic allele that produces a PEX1
protein with partial functionality and, as such, is generally associated with a milder phenotype.
Because it is common amongst ZSD patients and produces a PEX1 protein with some
functionality, this mutant is a major target for ZSD therapy development efforts. The ability to test
candidate bioactive small molecules identified in by compound library screening efforts on cells
from various tissue types would be of great value to validating these molecules and bringing them
to the clinical level more rapidly. However, we rely upon immortalized ZSD patient-derived
fibroblast models compound heterozygous for the PEX1 c.2528G>A p.G843D and p.I700fs
mutations. Although valuable, these cells may not accurately reflect the effect that our candidate
small molecules have on cells of affected tissue types, such as liver, retinal, and neural cells. One
goal of our lab is to produce such models bearing the PEX1 c.2528G>A p.G843D mutation to test
our small molecules. Because the PEX1 c.2528G>A p.G843D mutation is a single point mutation,
Cas9-base editor technology is a prime candidate for producing the PEX1 c.2528G>A p.G843D
mutation in different cell types.
The Cas9-base editor technology could also be used to correct the PEX1 c.2528G>A
p.G843D mutation by converting the mutant A base back to the wild type G. The rescue of a single
allele carrying the PEX1 c.2528G>A p.G843D mutation is enough to compensate for peroxisome
assembly defects in the presence of a second null or hypomorphic allele. Thus, the ability to correct

28
the PEX1 c.2528G>A p.G843D mutation on one allele could potentially be of significant
therapeutic value in the future.
The Liu laboratory at the Broad Institute at Harvard University has recently published on
their new Cas9-base editor technologies and have shown great promise for the technology in their
efforts. Currently, they have described two systems: 1) a nickase-Cas9 fused to a cytidine-
deaminase [74] and 2) a nickase-Cas9 fused to a adenosine-deaminase [75]. In the case of the
Cas9-cytodine-deaminase, the Liu lab has produced an engineered protein with rAPOBEC1 fused
at its N-terminus. rAPOBEC1 is an RNA cytidine-deaminase that can also act on DNA and
converts cytidine (C) to uracil (U), which shares base pairing properties with thymine (T)
(Figure8a). This effectively converts a C to a T permanently and inheritably. When targeting the
coding strand, the system can produce C to T mutations, while targeting the non-coding strand will
produce G to A transition mutations in the coding strand. Because rAPOBEC1 can only act on
single stranded DNA and rejects double stranded DNA, the base editor can act only in a narrow
region of about 8bp within the single stranded bubble created by the Cas9 in the targeted
protospacer sequence. This makes the system highly targetable and precise, and eliminates the
possibility of deaminating cytosine residues outside the targeted protospacer. In addition to this,
because it does not produce double strand breaks, the risk of producing indels at the target locus
and other off target loci is drastically reduced as compared to the traditional CRISPR-Cas9 system.
There are no naturally occurring adenosine-deaminase enzymes that can act on DNA, and
instead they only bind and convert adenosine residues on RNA strands, usually within tRNA.
Because of this, the Liu lab had to evolve an adenosine-deaminase capable of binding DNA. They
chose the E. coli adenosine-deaminase TadA as a starting candidate since it shares homology with
the APOBEC1 enzyme they had previously used for their cytidine-deaminase system. After

29
extensive testing of unbiased libraries of TadA-dCas9 fusion containing mutations within the
TadA portion of the protein, they were able to delineate that mutations at D108 eliminated the need
for TadA binding of the 2’-OH group of a ribose molecule, effectively reducing the energetic cost
of binding to DNA. All in all, they were able to produce a TadA-Cas9 (ABE) fusion protein that
can act on DNA (Figure 5b). Similar to the APOBEC1-Cas9 enzyme, this system can only act on
single-stranded DNA, so is only effective in the small bubble created by Cas9 upon binding to the
targeted region. It also can only produce nicks on a single strand of the DNA, so double-strand
cleavages do not occur. Again, this system is highly targetable, precise and less prone to indel
formation compared to traditional CRISPR-Cas9 systems.

30

Figure 5. Overview of base editing by Cas9 base editors. (A) Cytidine deaminase base editor. Diagram depicts
cytodine deaminase base editor (BE3) targeting an C residue in targeted genomic DNA. Like traditional CRISPR, a
PAM is required for Cas9 targeting and binding. (B) Adenosine deaminase base editor. Diagram of adenosine
deaminase base editor (ABE). The same basic principles apply for this system but instead it targets A residues. Note,
the editing window for both ABE and BE3 is in the sequence distal to the PAM, from about base 2 to 8. Note the base
editor only edits the strand that is not base-paired to the gRNA, thus the gRNA sequence should be identical to strand
that is to be edited. Figure taken from reference [74, 75].

Using these CRISPR base editing systems, we hypothesized that the PEX1 c.2528G>A
p.G843D mutation may potentially be a prime candidate for both the creation of the mutation
(BE3; G to A mutation) and the correction of the mutation (ABE; A to G mutation).

31
For the BE3 creation of the PEX1 c.2528G>A p.G843D mutation, we searched the
mutation loci for a suitable protospacer adjacent motif (PAM) with sequence NGG which is
required for Cas9:gRNA detection and binding. Because this mutation requires a G to A mutation
on the coding strand, it was necessary to find a PAM and protospacer on the anticodon strand in
order to target the corresponding C and convert it to a T, thus producing an A on the codon strand.
The only suitable protospacer-PAM sequence within the locus of interest placed the target base
just within the window of activity of the cytidine deaminase at position 3 distal to the PAM. The
optimal window as reported by Liu et al is within positions 2 through 8 distal to the PAM, with
decreasing activity closer to the extreme proximal end. Because our target base was close to the
extreme end, concern arose over the efficiency of conversion of the target base. In addition to this
concern, the protospacer also contained two C residues in positions 4 and 8, placing them well
within the activity window. To address this, we hypothesized that increasing the length of the
gRNA could help place the target base within a more optimal window of activity and potentially
push the C at position 8 out of the window. We designed the following four gRNAs, each differing
by the addition of a single bp (PAM denoted in brackets):
5’-CACCAATCTTGTCCCAACCC [AGG]-3’
5’-CCACCAATCTTGTCCCAACCC [AGG]-3’
5’-CCCACCAATCTTGTCCCAACCC [AGG]-3’
5’-ACCCACCAATCTTGTCCCAACCCC [AGG]-3’
To correct the PEX1 c.2528G>A p.G843D mutation, we searched for a PAM and
protospacer that targeted the coding strand since the intended conversion is from an A to G and
placed the target A base within the activity window of the Cas9-ABE fusion protein. Here, we
were able to find a suitable PAM that placed the target A in position 4 distal to the PAM well

32
within the optimal activity window. Luckily, the mutant A is the only A within the activity
window, so there was no concern for conversions of unintended bases. The following gRNA was
used for this purpose (PAM in denoted in brackets):
5’ TTGATGGGTTACATGAAGTT [AGG]-3’
Materials and Methods

gRNA Cloning

All gRNAs were cloned into the gRNA Cloning Vector (Church Lab) purchased from
Addgene. Following the protocol recommended by the Church lab, DNA oligos were designed
and purchased from Thermo Fisher Scientific. For each gRNA, the desired protospacer was
incorporated into the following forward and reverse oligonucleotides (undefined bases represent
where gRNA sequence is incorporated):
Insert_F:
5’-TTTCTTGGCTTTATATATCTTGTGGAAAGGACGAAACACCGNNNNNNNNNNNN
NNNNNNN-3’
Insert_R:
5’-GACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAACNNNNNNNNNNNNN
NNNNNNC-3’
Oligo pairs were annealed and incorporated into the linearized gRNA Cloning Vector after
digestion with AflII via Gibson Assembly. The resulting product was transformed into chemically
competent E. coli (NEB) and plated on kanamycin (50mg/mL) LB agar plates. Colonies were
picked the following day, mini-prepped with the GeneJet Plasmid Midiprep Kit (Thermo Fisher
Scientific) and sent for sequencing (Genewiz) to check for correct incorporation of gRNA annealed
oligo fragment.

33

Cell Culture

To introduce the PEX1 c.2528G>A p.G843D mutation using Cas9-BE3, we chose
unaltered HEK293T cells as a prime candidate cell type to transfect due to ease of transfection and
normal PEX1 gene. For Cas9-ABE correction of PEX1 c.2528G>A p.G843D mutation, we used
our immortalized PEX1 null/G843D GFP-PTS1 primary fibroblast cell line. All cells were cultured
at 37°C with 5% CO
2
with Fibroblast medium (high-glucose DMEM supplemented with 10% FBS,
penicillin/streptomycin, vitamin solution, essential and nonessential amino acids, all from Thermo
Fisher Scientific).

Plasmid Transfection

293T cells were co-transfected with 1ug of total DNA in 1:3 ratio of gRNA:Cas9-BE3
plasmid vectors on 48-well plates seeded at 60,000 cells/well. Transfections were performed one
day after seeding using Lipofectamine LTX (Thermo Fisher Scientific) with 2 uL Lipofectamine
LTX, 1 uL Plus reagent mixed with plasmids in 20 uL total OptiMem (Thermo Fisher Scientific)
following manufacturers recommended protocol.
For immortalized primary fibroblast transfections, cells were plated at 5X10^4 cells/well
the day before transfection onto 24-well plates. Each well was co-transfected with 0.5 ug total
DNA at 1:3 ratio of gRNA:Cas9-ABE. DNA mixed with 1 uL Lipofectamine LTX and 1 uL Plus
reagent in total of 40 uL OptiMem, following manufacturers protocol.
Immunofluorescence

34
To monitor primary fibroblast rescue of peroxisome assembly, we used
immunofluorescence imaging for GFP puncta. Monitoring for the presence of GFP puncta began
three days after transfection.
Cell isolation

To isolate GFP puncta positive primary fibroblasts, cells were first located via
immunofluorescence and position on plate marked. Media was removed and glass isolation rings
dipped in silicone grease were then placed around the marked area. Cells within the ring were
trypsinized with 0.05% trypsin (Thermo Fischer Scientific) and moved to 15-mL conical tube.
Total cells were then counted using a hemocytometer. Cell suspension was then diluted to achieve
1 cell/well and plated onto 96-well plates. Wells containing GFP puncta positive cells were then
identified 1-2 days after seeding and allowed to expand for downstream uses.
PCR and Sequencing

For detection of mutation creation in 293T cells, the following primers were used:
gPEX1_F14-1: 5’-GTCTATAGATTTGTCAACCTGATTTTC-3’
gPEX1_R15-2: 5’-AGCAGGCTCCTTAACCCAG-3’
For detection of mutation correction in primary fibroblasts, the following primers were
used, which capture the G843 loci as well as the I700 null frameshift loci:
hPEX1_e13F1: 5’-GGTCAACCTTAGAGTATC-3’
hPEX1_e15R: 5’-CTTCTGGGAGTAAGTATTCAC-3’
Genomic DNA was extracted from cells using the Purelink Genomic DNA Extraction Kit
(Thermo Fisher Scientific) according to manufacturer’s protocol. All PCRs were performed using
Phusion polymerase (Thermo Fisher Scientific) with 150 ng of extracted gDNA. For 293T PCRs,
35 cycles with Tm=62C and extension time of 1.25 minutes was used. For primary fibroblasts, 35

35
cycles with Tm=54C and extension time of 1.5 minutes was used. PCRs were purified using the
QIAquick PCR Purification Kit and sent for sequencing with the following primers:
hPex1seq_e14F1 5’-CACTATAGATTTGTCAACCTG-3’ (to sequence I700fs locus)
hPex1seq_e15F1 5’-GTTTCCAGCTAAGATGATGG-3’ (to sequence G843D locus)
Subcloning

PCR products were subcloned into the pCR Blunt II-TOPO vector (Thermo Fisher Scientific)
following manufacturers protocol and resulting products transformed into chemically competent
E. coli cells (NEB).

Results

Generation Of The PEX1 c.2528G>A p.G843D Mutation in HEK293T Cells

Liu et al report up to 70% base editing with the BE3 vector system in their experiments
[74]. Due to these prior observations, we reasoned that we should be able to detect the presence of
mutations caused by the base editor system in our 293T cells from the bulk population of
transfected cells. Indeed, upon sequencing PCR products from the bulk population, we saw
evidence of G to A conversions occurring within the protospacer of the targeted PEX1 loci.
To further characterize the mutations that had been achieved, we then subcloned the same
PCR product into E. coli and sequenced plasmids from the resulting process. Of the 109 individual
colonies picked and successfully sequenced, 25 had mutations in the target region, giving us an
estimated 23% overall efficiency in base editing. However, of these 25, only one had the intended
mutation alone, while others either contained one or two unintended mutations within the
protospacer (Figure 6). Overall, the efficiency of intended base conversion was only 0.92%.
Although small, this is still promising in that we demonstrated that producing the PEX1

36
c.2528G>A p.G843D allele in cells that do not originally possess it is possible. Additionally, we
did see a difference in the targeting of our intended base among the varying gRNA lengths used.
Of the four tested, the gRNA of length 22-bp was most successful at targeting our base. However,
it did not seem to affect the targeting window as the other two A residues at position 4 and 8 distal
to the PAM were still consistently converted at a higher rate than the intended base.

Figure 6. Results of BE3 PEX1 transfection in HEK293T cells (A). Sequencing chromatograms results from bulk
population of HEK293T transfected cells with gRNAs of lengths. All show evidence of successful base editing, except
gRNA 23-bp. Most robust editing occurred with gRNA of length 22-bp. (B.) PCR product from cells transfected with
BE3 and gRNA 22-bp were subcloned and 109 subclones were successfully sequenced. Of these, 25 subclones
contained base editor mediated mutations. Only one subclone showed the desired mutation, converting only glycine
843 to asparagine, while all other subclones contained undesired mutations.

Correction of PEX1c.2528G>A p.G843D in patient immortalized primary fibroblasts

After initial transfection, cells were monitored via immunofluorescence for the appearance
of clear GFP puncta. After five days post-transfection, small disparate patches of GFP puncta
positive cells became evident (Figure 7). Visual estimation approximates the percentage of rescue
somewhere around 1-5%, which is not very efficient but still a promising result. Because
appearance of puncta is not sufficient evidence for rescued peroxisome function via correction of

37
mutation, we isolated single cells with puncta by serial dilution. These cells were allowed to
expand until there were enough to collect for gDNA extraction (Figure 8).

Figure 7. Fluorescence imaging of PEX1 I700fs/G843D immortalized patient fibroblasts before and after
transfection with ABE-PEX1 gRNA. (Left). Untreated PEX1-p.G843D/I700fs immortalized patient fibroblasts.
No cells show puncta, indicative of impaired peroxisome assembly. This is representative of population. (Right).
Two days after transfection with ABE-PEX1gRNA plasmids, small, disperse populations of cells were seen with
bright GFP-PTS1 puncta, suggesting successful correction of the PEX1-p.G843D mutation back to its wild-type
state and the recovery of peroxisome assembly.

38

Figure 8. Isolated colony of ABE-PEX1 transfected immortalized fibroblasts showing rescue of peroxisome
assembly. Colonies with GFP puncta were isolated by serial dilution and plated on 96-well plates and allowed to
expand. Above is an example of one of the isolated cell colonies with apparent rescued peroxisome assembly after
five days of expansion. Note, all cells have robust GFP puncta, suggesting a pure population of genetically corrected
cells. This also shows that the corrected gene is passed on to daughter cells after mitosis.
To genotype these cells, we used primers that spanned the PEX1 I700fs null mutation site
and the PEX1-p.G843D mutation site of the PEX1 gene so that we could make sure the correction
was occurring on the correct haplotype. In other words, if we only amplified the PEX1-p.G843D
mutation site, we would not be able to tell if we were looking at the PEX1-p.I700fs allele, which
looks wild-type at the G843 loci. Upon subcloning and sequencing these PCR products, we found
evidence of alleles which did not contain either the PEX1-p.I700fs or the PEX1-p.G843D
mutation, suggesting the Cas9-ABE had successfully targeted the mutation and corrected it.
Surprisingly, we also found evidence of alleles that contained both the PEX1-p.I700fs and PEX1-
p.G843D mutations. Upon comparing this result to untreated primary fibroblasts cells, we found
evidence of the presence of the same double mutant allele. We reasoned that some sort of
homologous recombination event had occurred naturally in the cells to produce the double
mutation allele. However, this posed as a problem, as it is possible that a similar recombination

39
could have occurred to produce an allele with no mutation. We do not believe this to be the case,
however, since we do not find rescued GFP-puncta positive cells in our untreated fibroblast cells.
Though these results are modest in their efficiency, we are hopeful for the future
applications of this technology. Further research is required to examine the possibility of
increasing the efficiency of base editing via transfection. In addition, it may be more effective if
delivery of the Cas9-ABE system is implemented in alternative ways, including lentiviral and
AAV viral delivery. We will continue to explore the possibilities of this technology and implement
it to its furthest ability.

40
CHAPTER 5: FUTURE DIRECTIONS

HepG2 PEX1-null CRISPR-modified cell lines

To date, cell culture models of PBDs have been limited to primary and immortalized patient
fibroblasts. Although they have proven invaluable in disease diagnostics and also in the screening
libraries of small molecules, they may not adequately reflect the action of the small molecules in
specific tissue types. The HepG2 Pex1-null hepatocyte cell line models developed in our
laboratory provide potentially powerful new tools for investigating the underlying cellular
mechanisms of PBDs and evaluating the effectiveness of small molecules to rescue peroxisome
assembly in the context of liver-like cells. With these cells, we can further validate our small
molecule candidates by assessing their effectiveness in these hepatocyte-like cells.
We have already shared the HepG2 PEX1-null cells with colleagues working in the field
of peroxisome research, including Nancy Braverman (McGill University) and James Inglese
(NCATS/NIH). They have validated that the cells are peroxisome-deficient and have begun their
own independent research studies utilizing the cells. Additionally, we have begun testing some of
our more promising small molecules discoveries on these PEX1-null cells to assess whether
peroxisome function can be rescued. A drawback that may limit the usefulness of these cells in
small molecule discovery experiments is that they are PEX1 nulls with no residual activity. From
our laboratory’s experience with primary and immortalized patient fibroblasts, it would appear
that the rescue of peroxisome function is far more likely when a PEX1 protein with residual activity
is present (e.g. PEX1-p.G843D protein) than in cases where no PEX1 protein is present. Therefore,
we also hope to develop HepG2 cells that carry the PEX1-p.G843D mutation. Having cells that
are PEX1-p.G843D homozygotes, PEX1-p.G843D carriers, and compound heterozygous PEX1-
p.G843D/null cells would broaden the scope of our small molecule screens and may better reflect

41
the potential usefulness of our candidate small molecules. We hope to apply gene editing
technology to create HepG2 cells with the above genotypes.

BE3 PEX1 Editor (PEX1-p.G843D mutation generation)

Although our results with the BE3 cytidine deaminase Cas9 editor in converting the WT
Pex1 allele to one carrying a the PEX1-p.G843D mutation were modest, we are hopeful that we
can optimize the implementation of the technology to achieve more robust results. This may
include alternative ways of delivering the BE3-gRNA vector package, including delivery via
lentiviral and adeno-associated virus (AAV) infection or through the concurrent electroporation of
BE3 protein and PEX1 targeting synthetic gRNA. To date, we have only been able to test
transfection with this system and do not know how effective these other forms of delivery may be.
We hope to explore these possibilities in the near future.
In addition, there are numerous other immortalized cell lines where we could, in principle,
introduce mutations in PEX genes, including the PEX1-p.G843D allele. We plan to use the BE3
system to introduce the PEX1-p.G843D mutation into more cell lines representative of various
affected tissue types. Having cell lines of different lineages carrying the PEX1-p.G843D mutation
would be of much importance to our research as it would allow for experimentation of disease
mechanism in different cell types and give us the opportunity to try out our small molecule
candidates on various cell types. Again, this would help us further validate candidate small
molecules and increase our confidence that they would be effective at treating the disease in the
most heavily affected tissue types.
The key technical issue involves determining the copy number of the PEX gene in question
that is to be genetically altered. It is well-known many immortalized cells lines are aneuploid. As

42
such, we would need to determine the copy number of the PEX gene in question first using
cytogenetic techniques including G-banding, fluorescence in situ hybridization (FISH) and array
cytogenetic hybridization (aCGH). In cases where the PEX1 gene is present in expanded copy
number, one would need to mutate multiple copies of the gene to achieve a phenotype. Ideally one
would introduce a single PEX1-p.G843D allele in a background of all other alleles being PEX1
null. It is much more simple in the case of cell lines with just one copy of the PEX gene, since only
one copy of needs to be mutated. One would predict that it would be highly efficient to generate
such cell lines due to the need to genetically alter just one chromosome.

ABE Pex1 Editor (PEX1- p.G843D mutation correction)

Results with the ABE PEX1 gRNA editing to correct the PEX1-p.G843D mutation were,
again, modest at best. However, we again believe this to be an issue of providing sufficient DNA
delivery to the cells and not an issue of the performance of the system. We are exploring the
possibility of using different delivery methods such as lentiviral and AAV viral delivery, to
introduce the base editor system to our cells. This would allow for more efficient delivery and also
prolonged expression of the base-editing transgene.

In vivo base editing

If viral delivery is an option soon, we may be able to adapt the system to target the Pex1-
p.G844D mutation in mouse, the orthologous mutation to the human PEX1-p.G843D mutation in
humans. The homozygous Pex1-p.G844D mouse model of mild ZSD, co-developed by Drs. Nancy
Braverman (McGill University) and Steven Steinberg (Kennedy Krieger Institute), was first
reported in 2014 [76]. The homozygous Pex1-p.G844D mutant mice showed classic phenotypes

43
associated with mild ZSD including growth retardation (Figure 12), progressive retinopathy, and
fatty liver disease (Figure 13). Importantly, there was robust biochemical and histological evidence
of a bile acid defect that is associated with intestinal fat malabsorption and cholestasis in the young
homozygous Pex1-p.G844D mice [76]. These homozygous Pex1-mutant mice show elevated
blood VLCFA levels relative to controls (10.7-fold, P<1×10
-12
), indicative of impaired
peroxisomal lipid catabolism [76]. They also show reduced blood plasmalogen levels relative to
controls (2.1-fold, P<1×10
-4
), indicative of impaired peroxisomal biosynthetic pathways [76]. This
mouse model would provide a powerful opportunity for testing out the ABE-mediated correction
of the murine equivalent of the PEX1-p.G844D mutation in vivo and thus allow us to test the
efficacy of the system in living animals. This would be an invaluable opportunity and would
immensely benefit the push towards translating this technology to a clinical application. We are
excited and hopeful for the future of this technology.

Figure 12. Growth patterns of Pex1-p.G844D homozygotes from day 7 to 98 postnatal. Panels A and B show growth
curves male and female Pex1-p.G844D homozygotes from 7 to 21 days postnatal. Grey represents Pex1-p.G844D mice while
blue represents wild-type mice. Panel C shows side by side comparison of size of Pex1-p.G844D heterozygote (left) and Pex1-
p.G844D homozygote (right) litter mates. Note the significant reduction in growth in the homozygote. D shows growth in both
sexes from days 28 to 98 as compared to wild-type mice. Figure taken from reference [76].

44

Figure 13. Liver histologies of G844D homozygous mice. (A) WT control mouse liver histology. (B) and (C) Histological
analyses of two homozygous Pex1-p.G844D mouse livers. Cytoplasmic volume is increased in G844D homozygotes as
compared to control characteristic of microvesicular fat deposition. Small arrows indicate yellowish cholestatic deposits not
present in control liver. Large arrow in C shows bile ductular cell proliferation, suggesting a response to bile duct damage in
the area. Figure taken from reference [76].

45
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Abstract (if available)

Abstract Peroxisomes are microbody organelles present in virtually all cells of eukaryotic organisms. They play vital roles in numerous metabolic pathways including the catabolism of very long chain fatty acids (VLCFAs) as well as the biogenesis of docosahexaenoic acid (DHA) and plasmalogens. Proper assembly and function of peroxisomes is essential to human health and development. The importance of proper peroxisome function is highlighted by a number of multi-systemic disorders in humans that result from inherited mutations in peroxisome-related genes, collectively referred to as peroxisome biogenesis disorders (PBDs). Peroxisome biogenesis disorders affect the body globally, although certain organ systems that rely more heavily on peroxisome-mediated metabolism are more seriously affected, including the nervous and hepatic systems. ❧ Currently, because primary cell models are inaccessible due to ethical dilemmas, it is difficult to study the mechanics of PBDs in the context of specific organ systems. The most common cell-based models for PBDs are derived from patient and genetically engineered mouse fibroblasts. Although these cells have proven invaluable for screening small molecule libraries, our goal is to establish and test cell models that more closely represent affected cell-types. We hypothesized that the widely utilized HepG2 liver-cancer immortalized cell-line carrying common PBD mutations may serve as a better model for testing the effects that drugs identified as possibly therapeutic in immortalized fibroblast models may have on liver function and health and better elucidate the mechanisms by which the rescue is occurring. We used CRISPR/Cas9 gene editing to introduce PEX1 null mutations in HepG2 cells via the introduction of double-strand breaks in PEX1. We were able to successfully generate two separate HepG2 cell lines, each homozygous for null PEX1, differing slightly in their specific mutations. ❧ We also explored the use of a new CRISPR-Cas9 base editing system, developed in David Liu’s laboratory at Harvard University, for its ability to both introduce and correct the most common PEX1 mutation, the PEX1-p.G843D mutation. This single-base pair mutation results in a hypomorph allele with limited gene function. Using the CRISPR-Cas9 base-editing system, we have preliminary evidence that suggests both introduction and correction of PEX1-p.G843D are possible and efficient. Further research is required to investigate the potential benefits to PBD research and therapeutic endeavors that this technology may make possible.

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Creator Steele, Bradford Harrison (author)

Core Title Developing novel in vitro model systems to investigate therapeutic hypotheses for peroxisome biogenesis disorders

School Keck School of Medicine

Degree Master of Science

Degree Program Biochemistry and Molecular Biology

Publication Date 05/10/2019

Defense Date 03/14/2019

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

Tag base editor,CRISPR-Cas9,OAI-PMH Harvest,peroxisome biogenesis disorders,peroxisomes,Zellweger syndrome

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Hacia, Joseph (committee chair), Frenkel, Baruch (committee member), Stallcup, Michael (committee member)

Creator Email bhsteele@usc.edu,bradfordhsteele@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c89-157601

Unique identifier UC11660967

Identifier etd-SteeleBrad-7433.pdf (filename),usctheses-c89-157601 (legacy record id)

Legacy Identifier etd-SteeleBrad-7433.pdf

Dmrecord 157601

Document Type Thesis

Format application/pdf (imt)

Rights Steele, Bradford Harrison

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA