LI LIU Master Thesis 111316

Home , Li (surname 李)

TARGETED MUTAGENESIS OF ZEBRAFISH HEARING-RELATED

GENES USING ZFN AND TALEN

LI LIU

Submitted in partial fulfillment of the requirements

For the degree of Master of Science

Thesis Advisor: Dr. Brian M. McDermott

Department of Biology

CASE WESTERN RESERVE UNIVERSITY

January, 2014

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Li Liu ______

Master of Science candidate for the ______degree *.

Jean H. Burns (signed)______(chair of the committee)

Brian M. McDermott ______

Claudia Mizutani ______

Emmitt R. Jolly ______

______

10/28/2013 (date) ______

*We also certify that written approval has been obtained for any proprietary material contained therein

TABLE OF CONTENTS

LIST OF FIGURES...... 2 ACKNOWLEDGEMENT...... 3 LIST OF ABREVIATIONS...... 4 ABSTRACT...... 5 INTRODUCTION...... 6 Hearing and hair cells...... 6 Hearing related genes...... 11 Zebrafish as a model organism...... 15 Gene targeting technology...... 16 MATERIALS AND METHODS...... 21 RESULTS...... 28 CoDA-ZFN induced somatic mutations in zebrafish fascin 2a gene in a site-directed manner...... 28 CoDA-ZFN induced site-specific heritable mutations in zebrafish piezo 2-4 gene...... 32

TALEN induced heritable mutations in zebrafish piezo 2-4

gene with a higher efficiency...... 38

DISCUSSION...... 43 REFERENCES...... 48

LIST OF FIGURES

FIGURE 1...... 7 FIGURE 2...... 8 FIGURE 3...... 10 FIGURE 4...... 12 FIGURE 5...... 14 FIGURE 6...... 17 FIGURE 7...... 18 FIGURE 8...... 20 FIGURE 9...... 27 FIGURE 10...... 29 FIGURE 11...... 30 FIGURE 12...... 31 FIGURE 13...... 32 FIGURE 14...... 33 FIGURE 15...... 35 FIGURE 16...... 35 FIGURE 17...... 37 FIGURE 18...... 39 FIGURE 19...... 40 FIGURE 20...... 41 FIGURE 21...... 42 FIGURE 22...... 44 TABLE 1...... 22 TABLE 2...... 22 TABLE 3...... 30 TABLE 4...... 31 TABLE 5...... 34 TABLE 6...... 39

ACKNOWLEDGEMENT

My first acknowledgement and deepest appreciation goes to my advisor Dr.

Brian M. McDermott for his excellent guidance, patience and caring. He encouraged me to become not only an experimentalist, but also an independent thinker on research. He spent time answering my questions, as well as encouraging and motivating me to find out the answer on my own. He has been a great mentor as well as a thoughtful friend who cares about my life, my family and supports my long-term objectives.

I also would like to express gratitude to the rest of my committee members, Dr.

Claudia Mizutani and Dr. Emmitt R. Jolly for their encouragement, insightful comments, and hard questions. I greatly appreciate their time talking to me and responding to my questions.

I would like to thank Shih-wei Chou, who as a good friend and a co-worker has been always willing to instruct and help since the first day I worked with her. Many thanks to Carol Fernando, Jiaqi Hu, and other people in the laboratory for supporting me in work and in daily life. My research would not been possible without them.

I would like to thank the Department of Biology at Case Western Reserve

University for the diverse resources and help they provided.

Finally, my heartfelt appreciation goes to my parents, who have been supporting me psychologically and financially in the past 23 years. Their continuous love provided me courage to pursue my research at the other end of the world.

LIST OF ABBREVIATIONS

NIDCD, national institute of deafness and other communication disorder; F-actin, filamentous actin; ZFN, zinc finger nuclease; TALEN, transcription activator-like nuclease; FSCN, fascin; PKC, protein kinase C; MA, mechanically activated; Dmpiezo,

Drosophila Piezo; Mmpiezo, mouse Piezo; RNAi, RNA interference; MOs, morpholino oligos; ZF, zinc finger; NHEJ, non-homologous end joining; CoDA, context-dependent assembly; TAL, transcription activator-like; RVD, repeat variable di-residues; HRMA, high melting resolution analysis; dpf, day-post-fertilization; Tm, annealing temperature; sgRNAs, single guide RNAs

Targeted mutagenesis of zebrafish hearing-related genes using ZFN and TALEN

Abstract

LI LIU

Hair cells are specialized mechanosensory cells with stereocilia protruding from their apical surfaces. Deflection of stereocilia allows the opening of mechanotransduction channels on the tips of the stereocilia. However, regulation of stereocilia development at the molecular level and the identity of the mechanotransduction channel remain largely unknown. Engineered ZFN and TALEN have shown to be efficient genome editing tools among many different organisms. We designed ZFN or TALEN targeting two of our genes of interest: fascin 2a and piezo 2-4 in zebrafish. The ZFNs and TALENs were constructed and successfully induced somatic mutations at the targeted loci for both genes. Founder fish that carry and pass mutation to their offspring were also found for the piezo 2-4. With the success in introducing insertion/deletion mutations within the desired target loci, we are now able to investigate the function of a gene in hair cell genesis and mechanotransduction in zebrafish.

Introduction

Hearing and hair cells

Hearing is one of our primary senses that serve important functions for assisting with individual safety and for communicating with the outside world. Hearing loss can increase susceptibility to physically harmful situations, restrict one’s ability to effectively interact with others, and even may lead to feelings of isolation and depression. Statistics from the National Institute of Deafness and Other Communication

Disorder suggest that approximately 17% (36 million) of American adults report some degree of hearing loss.

The ear is the organ primarily responsible for hearing. It functions as a transducer, translating external sound waves into electrical signals that can be interpreted by the brain. The mammalian ear is composed of the outer, middle, and the inner ear (Figure 1A). The outer ear collects sounds from the external environment and channels them to the middle ear. Sound waves hit the tympanic membrane in the middle ear, causing the vibration of the tympanic membrane that vibrates the bones of the middle ear. The inner ear has two structures: the vestibular labyrinth and cochlea, which function in balance and hearing, respectively. The hair cells localized to the cochlea are shown in Figure 1B. When vibration from the middle ear is transferred into the inner ear cochlea, the basilar membrane oscillates, causing hair cells to move back and forth relative to the position of the tectorial membrane. This shearing motion causes the depolarization of hair cells. Hence, the mechanical movement gets translated into electrical signal and further propagates into the brain.

Figure 1. Mammalian ear structure. (A)The mammalian ear has three components: external, middle, and inner ear. The external ear collects sounds from outside environments. The middle ear ossicles vibrate in response to sound waves transmitted into the middle ear. The inner ear consists of the vestibular labyrinth for balance and the cochlea for hearing. (B) Cross-section of the inner ear cochlea reveals the organ of Corti. It consists of three rows of outer hair cells and one row of inner hair cells sitting between the basilar membrane and the tectorial membrane. Adapted from Frolenkov et al., 2004. Originally published in Nature Review of Genetics. 5:489- 498. Hair cells are the mechanosensory receptors in both the vestibular labyrinth and cochlea of the inner ear. The importance of hair cells is further emphasized by the statistics from NIDCD, which state that the predominant cause of hearing loss is impairment of hair cells. Therefore, the molecular mechanisms underlining hair cell morphology and function are important for the study of deafness.

Hair cells obtain their name from a hair-like bundle protruding from their apical surface, called the hair bundle (Goodyear et al., 2006). The hair bundle is composed of actin-based stereocilia arranged as a staircase shape and a single microtubule-based kinocilium that is next to the tallest row of stereocilia; however the kinocilium is degenerated in mature mammalian cochlear hair cells (Figure 2) (Vollrath et al., 2007).

Figure 2. Schematic diagram of a hair cell flanked by supporting cells. At the apical surface of the hair cell is the hair bundle, while nerve endings contact the hair cell along its basolateral region. The hair bundle is composed of stereocilia with increasing length and a single kinocilium adjacent to the tallest stereocilia. Stereocilia are extended to the cuticular plate, an actin-enriched meshwork on the apical cytoplasm of the hair cell. Adapted from Kandel, Schwartz & Jessell. Originally published in Principles of Neural Science, 4th Edition. Stereocilia are filled with numerous, highly cross-linked filamentous actin (F- actin) (Figure 3). Actin-binding and -bundling proteins, through interactions with F- actin, serve a variety of functions in structuring stereocilia (Nayak et al., 2007).

Dysfunction of these actin-interacting proteins may result in deformed stereocilia and sequentially lead to hearing loss. For example, espin proteins, that cross-link actin filaments, are responsible for lengthening and thickening stereocilia (Rzadzinska et al.,

2005). Moreover, myosin15a is essential for the regulation of elongation at the tip of stereocilia (Drummond et al., 2011). Mutations in either of these proteins cause 8 recessive nonsyndromic deafness in humans (Naz et al., 2004; Wang et al., 1998).

Each stereocilium is linked by four types of links externally: tip-links, top links, side-links and ankle-links. Tip links that connect the tip of one stereocilium to the side of the adjacent, taller stereocilia are thought to be important for mechanotransduction

(Pickles et al., 1984). Positive deflection of the hair bundle towards its tallest row increases tip link tension, resulting in the opening of ion channels therefore allowing some positive ions such as K+, and Ca2+ to flow into the hair cell (Lumpkin et al., 1995).

This process depolarizes the cell membrane of the hair cell and modulates the neurotransmitter release at its basolateral surface, thus triggering action potentials in the auditory nerve. The electrical signals that carry auditory and vestibular information are then transduced to the brain (Vollrath et al., 2007; Fettiplace, 2009).

Figure 3. Schematic diagram of two adjacent stereocilia. Stererocilia are composed of parallel actin filaments. Actin-bundling proteins are essential for the formation and maintenance of stereocilia. Adjacent stereocilia are linked by four types of links. Tip links are important for mechanotransduction. Tip links are anchored in the side-insertion plaque and tip-insertion plaque. Mechanotransduction channels sit on the tip of shorter stereocilia and can be opened by tip link tension. The other three types of links are top links, side-links and ankle-links, which are located at the base of stereocilia as represented in the figure. Electron microscopy reveals two electron dense regions on the stereocilia

(Figure 3). Mechanotransduction channels are thought to reside in these regions (Denk et al., 1995). However, a recent study which imaged the Ca2+ influx in rat hair cells

(Beurg et al., 2009) provided evidence that the channels are present only at the bottom of the tip links (Figure 3). So far, researchers have attempted to uncover the properties and identities of the mechanotransduction apparatus, in addition to determining their

10 localization in the hair bundle. Some of the actin-associated proteins and scaffolding proteins were found to be important to the function of mechanotransduction; however, little is known about the identity of the mechanotransduction channel itself.

Hearing related genes

Genes that are required for stereocilia formation and mechanotransduction are thus of great interest. In this study, our major focus is on fascin 2a and the piezo protein family members.

Fascins are actin-binding proteins which crosslink neighboring actin filaments into compact parallel bundles in diverse sub-cellular structures including the specialized actin bundles of photoreceptors and hair cells (Hashimoto et al., 2011).

Fascin was first discovered in the 1970s from purified cytoplasmic extracts of sea urchin (Bryan and Kane, 1978). Fascin 2 and fascin 3 were later identified in retina and testis, termed as retinal fascin and testis fascin respectively. Vertebrate fascin family members are highly conserved in amino acid sequences (Figure 4) (Kureishy et al.,

2002; Hashimoto et al., 2011). Homo sapiens fascin 1 monomer with 493 amino residues folds into four β-trefoil domains (Sedeh et al., 2010). Fascin 1 in the vertebrate fascin family binds and bundles F-actin to form a tightly packed parallel filamentous bundles, and this process is impaired upon phosphorylation by protein kinase C (PKC) at amino residue serine 39 (Yamakita et al., 1996; Adams et al., 1999). There are two predicted actin-binding sites in fascin 1 protein, and the bundling activity of fascin 1 was largely decreased when mutations occur in those two sites (Jansen et al., 2011).

Figure 4. Vertebrate fascin family members. Fascins are composed of four β-trefoil domains. The predicted actin-binding sites for fascin 1 are as shown. Red boxes represent the conserved PKC phosphorylation site. Adapted from Hashimoto et al., 2011. Originally published in The Journal of Pathology. 224:289-300. Fascin 2 is expressed in the inner and outer segments of retina photoreceptor cells. Similar to fascin 1, fascin 2 cross-links F-actin into compact bundles (Saishin et al.,

2000). Fascin 2 was recently found expressed in hair-cell stereocilia of frog, chicken, and mouse, and was shown to be essential for maintaining stereocilia length in mouse

(Shin et al., 2010). Fascin 2 mutant mice display progressive hearing loss in the high- frequency range (Perrin et al., 2013). Zebrafish have two fascin 2 paralogus: fascin 2a and fascin 2b. Fascin 2b has been shown to localize to hair-cell stereocilia and, when overexpressed, lengthen stereocilia (Chou et al., 2011). Fascin 2a, which shares 87% amino acid sequence similarity with fascin 2b, therefore is a candidate as a crucial actin-binding protein in hair cell stereocilia. Although the expression level of fascin 2a mRNA is relatively low in zebrafish hair cells (Chou et al., 2011), we can’t rule out the possibility that fascin 2a is upregulated to compensate for the loss of fascin 2b in hair

12 cells. Therefore, we will generate fascin 2a knockout zebrafish to further understand the role of the fascin protein family in shaping the morphology of stereocilia in hair cells.

Piezo proteins were first identified by Coste and collagues from the Neuro2A mouse neuroblastoma cell line in 2010. They are large transmembrane proteins conserved among various species (Coste et al., 2010). Vertebrates have two members in this protein family: piezo 1 and piezo 2. The predicted proteins contain more than 30 transmembrane domains as shown in Figure 5A and 5B (Coste et al., 2010). Similar to its mammalian counterpart, the drosophila piezo gene is also predicted to consist of a large number of transmembrane domains as shown in Figure 5C (Kim et al., 2012).

Besides structural analysis of piezo proteins, functional studies on the piezo family provided more evidence suggesting their crucial role in mechanotransduction.

Piezo 1 is required for mechanically activated (MA) currents in Neuro2A cells and piezo

2 is required for a subset of MA currents in dorsal root ganglion neurons (Coste et al.,

2010). Later, the same group showed that overexpression of drosophila piezo

(Dmpiezo) and mouse piezo 1 (Mmpiezo 1) in human cell lines gave rise to mechanically activated channels with distinct properties, suggesting that piezo proteins are channels responsible for cells’ mechanically gated response. The protein-interaction characteristics of purified Dmpiezo and Mmpiezo 1 also suggest that they themselves are channels but not scaffold proteins that collaborate with other channel proteins to function (Coste et al., 2012). In addition, Kim et al. demonstrated that in the fly piezo is essential for sensing noxious mechanical stimuli in vivo (Kim et al., 2012). From these studies in different species, I hypothesize that piezo proteins may play an important

13 role in mechanotransduction in zebrafish hair cells.

Figure 5. Transmembrane domains in mammalian piezo proteins and Drosophila piezo predicted by Transmembrane Hidden Markov Model (TMHMM) prediction program. (A) Hydrophobicity analysis of mouse Piezo1 shows succession of hydrophobic and hydrophilic regions. 30 transmembrane domains are predicted by TMHMM2 program analysis. Picture adapted from Coste et al., 2010. Originally published in Science. 330: 55-60. (B) Using the same analysis method with (A), 34 transmembrane domains are predicted in mouse Piezo 2. Picture adapted from Coste et al., 2010. Originally published in Science. 330: 55-60. (C) Dmpiezo is predicted to consist of 39 transmembrane domains as predicted by The Kyte-Doolittle hydropathy plot and TMHMM 2.0. Picture adapted from Kim et al, 2012. Originally published in Nature. 483:209-212. In this study, I provide a systematic method to study the function of candidate genes in hair cell development via creating gene specific knockout zebrafish.

Zebrafish as a model organism

Zebrafish (Danio rerio) has emerged as an excellent vertebrate model organism in hearing-related studies as well as in other research fields. There are numerous advantages of using zebrafish. Large number of rapidly developing and externally fertilized embryos as well as its transparency during embryonic stages made zebrafish an irreplaceable model for observing developmental process in vivo.

Another important advantage of zebrafish is that their hair cells are morphologically and functionally conserved with chick, mouse and human hair cells.

Several human deafness genes are also essential for zebrafish hearing and balance, further supporting the use of zebrafish for human hearing studies (Whitfield, 2002). In addition to hair cells found in the inner ear, zebrafish have hair cells found in the lateral line system along the body surface. This easy accessibility yields an additional advantage for hair cell research.

The use of the zebrafish as a research model organism has increased dramatically in the past 20 years, and the zebrafish genome-sequencing project started in 2001 has provided zebrafish researchers with helpful resources (Broughton et al.,

2001). There have also been a number of techniques developed to alter gene expression in zebrafish, including zebrafish transgenesis, forward genetics screening, reverse genetics tools such as Morpholino Oligonucleotides (MOs), Zinc Finger Nuclease (ZFN), and Transcription Activator-like Effector Nucleases (TALEN).

Gene targeting technology

Together with all the advantages in regard to genetic manipulations, the ability to generate site-directed gene modifications further increases the potential of this model organism. However, developing a knockout tool for targeting a specific zebrafish gene has been considerably challenging.

If we looked back at the history of gene targeting (Figure 6), the first knockout mice were generated in 1989 using embryonic stem cells (Koller et al., 1989). However, embryonic-stem-cell-based gene targeting has thus far not been successful in zebrafish

(Fan et al., 2006; Yan et al., 2009). RNA interference (RNAi) was first discovered in

Caenorhabditis elegans. It was demonstrated that exogenous introduction of double stranded RNA could trigger degradation of mRNA (Fire et al., 1998). However, RNAi has not been a reliable tool for zebrafish (Wargelius et al., 1999), since small interfering

RNA (siRNA) treatment led to inhibition of the microRNA (miRNA) pathway in zebrafish embryos (Zhao et al., 2008). Another knockdown technique, MOs, which work through an anti-sense mechanism, has been widely used (Bill et al., 2009). However, just like other knockdown techniques, MOs have drawbacks such as non-specific effects and incomplete and irreproducible gene silencing, which largely limits the use of this technique.

Figure 6. The history of gene targeting in zebrafish. Forward genetic studies have been carried out since late 1970s. However, even though knockdown technologies started to be applied to zebrafish in the late 1990s, site-directed knockout zebrafish were difficult to generate until ZFN technology and TALEN technology became available in 2008 and 2011, respectively. This was a large delay compared to the first knockout mouse generated in 1989. In 2008, Meng et al. and Doyon et al. generated site-directed mutations in zebrafish by injecting zinc-finger nucleases (ZFNs) into fertilized embryos (Meng et al.,

2008; Doyon et al., 2008), and demonstrated that ZFNs are capable to knockout a specific gene of interest in zebrafish. ZFNs are artificial restriction enzymes generated by fusing a DNA-binding domain with a FokI nuclease domain. The DNA binding domain is an array of Cys2His2 zinc finger proteins. Cys2His2 zinc finger is a group of zinc finger

(ZF) proteins that contain two histidine and two cysteine residues that hold a zinc ion in its approximately 30 amino acid motif (Brown et al., 1985). Each of them can be designed and engineered to recognize a 3 base pair target DNA (Hurt et al., 2003).

Dimerization of the FokI cleavage domain would efficiently create a double strand break in DNA (Mani et al., 2005). Repair of the double strand break by non-homologous end joining (NHEJ) introduces insertion or deletion mutations at the target site (Figure 7A).

ZFNs have been shown to induce mutations in human cells, drosophila, zebrafish, and other organisms (Gaj et al., 2012; Foley et al., 2009; Meng et al., 2008; Beumer et al.,

2008).

Since the sequence-recognition specificity of a given zinc finger in vivo is dependent on the context of its neighboring ZFNs, most ZFNs construction platforms which rely on in vitro selection have either low efficiency or high labor requirements

(Ramirez et al., 2008; Maeder et al., 2008). Context-dependent assembly (CoDA) is a rapid-to-use platform for fast assembly of engineered ZFNs, introduced by Sander et al. in 2011. With the CoDA approach, three-finger ZFN arrays are assembled using N- and

C-terminal fingers that have been previously identified in other arrays sharing a common middle finger (F2 units) (Figure 7B). Compared to previous platforms, CoDA is easy to generate, and is efficient. More importantly, ZFNs generated by CoDA are shown to induce mutations in the target site in zebrafish with success rates of 0.5% to

16.7% (Sander et al., 2011).

Figure 7. Schematic diagram of Zinc Finger Nuclease (ZFN) technology. (A) Engineered ZFN consists of a zinc-finger DNA binding domain, in which three zinc-finger arrays binds to 9bp of DNA sequence, and a FokI DNA cleavage domain. Two ZFNs function as a dimer, catalyzing double strand DNA breaks at the spacer region. Adapted from Kim et al., 2011. Originally published in Plant Biotechnology Reports. 5:9-17. (B) Schematic overview of CoDA. ZFN arrays for a target gene site can be created by joining two existing arrays, which have a common F2 unit. Adapted from Sander et al., 2011. Originally published in Nature Methods. 8:67-69. ZFN has been a crucial technique for zebrafish researchers with its ability of site- directed genome targeting. However, the context-dependent characteristic of ZFN and the availability of ZFN arrays for CoDA hugely limit the number of sites that can be targeted. This problem was then solved by a more recent technology: transcription activator-like effector nucleases (TALEN) in 2011 (Sander et al., 2011).

Transcription activator-like effector nucleases are also artificial restriction enzymes generated by fusing a DNA binding domain with a FokI DNA cleavage domain.

Different from ZFNs, the DNA binding domain of a TALEN is an array of TAL effectors.

Transcription activator-like (TAL) effectors are secreted by bacteria of the genus

Xanthomonas that cause diseases in plants such as rice and cotton (Boch et al., 2010).

One TAL effector binds to a single DNA nucleotide, and the recognition specificity is governed by two amino acids, also known as repeat variable di-residues (RVDs) (Boch et al., 2010) (Figure 8). Through proper construction of the TAL array, any sequence in the genome can be targeted by TALEN (Bogdanove et al., 2010). TALENs induced endogenous gene mutations in cultured human cell lines, yeast, zebrafish and other organisms with a comparable or higher success rate than ZFNs (Li et al., 2011; Miller et al., 2011; Cremak et al., 2011; Sander et al., 2011; Moore et al., 2012). The off-target effects of TALENs are reported to be rare. (Huang et al., 2011)

Figure 8. Genome targeting by TALEN. (A) Engineered TALE binds to a specific base. The recognition mechanism involves two key amino acid residues in 33-35 amino acid repeat. In the TALE shown as red, two key amino acid residues, HD, recognize base C. (B) Each TALEN consists of an array of TALEs forming the DNA binding domain and a FokI DNA cleavage domain. The dimerization of two TALENs triggers a double strand break in the target site region. DNA sequences that are recognized and bound by TALE are highlighted in red. Adapted from Clark et al., 2011. Originally published in Zebrafish. 8:147-149. Our aim in the following study is to establish a platform for generating knockout zebrafish by using ZFN and TALEN technologies. Using these techniques, answers to questions pertaining to the molecular basis of hair cell development and functional mechanotransduction might then be reachable. We designed and generated ZFNs or

TALENs for targeting multiple genes. By injecting ZFN- or TALEN-encoding RNA into zebrafish embryos, we successfully introduced heritable site-directed mutations in fascin 2a, piezo 2-4 and other hearing related genes.

Materials and methods

Zebrafish husbandry

Wild type Tübingen (TÜ) zebrafish (Danio rerio) were used to obtain embryos.

Zebrafish were bred and maintained according to the fish protocol (Westerfield, 2000).

ZFN and TALEN target site design

Gene sequences were obtained from the Ensembl database (Zv9

(GCA_000002035.2)). Sequences of exons were put into the ZiFiT Targeter to identify potential ZFN or TALEN target sites (Sander et al., 2011). Specificity of individual target site was tested by the NCBI BLAST tool. One pair of high resolution melting analysis

(HRMA) primers and one pair of sequencing primers were designed for each target site.

HRMA primer pairs met the following criteria: 1) amplify DNA product with length between 100 bp to 160 bp; 2) primer sequences are specific in the genome; 3) efficiently amplify the desired product after 25 cycles of PCR. Sequencing primer pairs were designed to amplify one single DNA fragment at 300-800 bp long.

Polymerase chain reaction (PCR)

PCR was performed using Ex Taq DNA Polymerase (Takara Bio) to amplify the target site with either HRMA or sequencing primer pairs. Genomic DNA isolated from 4 day-post-fertilization (dpf) zebrafish embryos was used as template in all PCR. PCR cycling conditions were as follows: pre-denaturation at 94°C for 2 min, followed by 35 cycles (25 cycles for HRMA primers) of amplification: 94°C for 20 s, 65°C for 45 s, 72°C for 30 s. Then allow the final extension at 72°C for 5 min. Annealing temperature (Tm)

21 may range from 65°C to 68°C for different primer sets as listed in Table 1 and Table 2.

Table 1. A list of sequencing primers used in this study. Sequencing primers were used for amplifying the target site region. Primer sequences, annealing temperature and the predicted product size are shown as below. Primer name Primer sequence (5’-3’) Tm (°C) Product

size (bp) fascin2a ZFN6-1 F GACGTTACCTGTCGTCCGAT 65 595 fascin2a ZFN6-1 R TGGAAAACGACTTGTGGGTGGCT zf piezo2-4 ZFN6-1 5’seq GAGGAAGTGGATGAGAACGACC 65 414 zf piezo2-4 ZFN6-1 3’seq2 GGATGCCATCCTCAAACTGGCTG

Piezo24 TALEN1 5’seq AAAACGACCTGCGTCCTGAG 65 564 piezo24 TALEN1 3’seq TGAAGTCAAATCTGGCCAAGAC

Table 2. A list of primers used for HRMA. Below is a list of primers we designed for HRMA analysis. Respective annealing temperatures and product sizes are shown as below. Primer name Primer sequence (5’-3’) Tm (°C) Product

size (bp) f2aZFN6-1 HRM F1 ACCTGTCCACCCAAGAAGATGAG 65 128 f2aZFN6-1 HRM R1 CTTGAGAAAGCGGTTGTCACAGG piezo24ZFN61 HRM F1 GCAAGCTGGTGATGGCTCTGCT 68 129 piezo24ZFN61 HRM R1 CGCAGGACAGCAGCATCATCA piezo24 TALEN1 HRM F1 ATGTCAGCTGGCTGACGTTTGTC 65 119 piezo24 TALEN1 HRM R1 CAGGAAGTTCCCATAAGCCACC

Construction of ZFN expression vectors

ZFN expression vectors are generated according to Forley et al., 2009. DNA

22 fragments that encode engineered zinc fingers were synthetically ordered from IDT

(Integrated DNA Technologies) as pIDTsmart-ZF. Heterodimeric ZFN expression vectors were made through subcloning the DNA fragment that digested from pIDTsmart-ZF vector into pMLM290 or pMLM292 vectors using XbaI and BamHI (New

England Biolabs). Colonies with expression vectors carrying either half site were selected and confirmed by sequencing with CMV_fwd_primer (5’-

CGCAAATGGGCGGTAGGCGTG-3’) and BGH rev_primer (5’-TAGAAGGCACAGTCGAGG-3’).

Construction of TALE repeats

Joung’s Lab TAL Effector Engineering Reagents and TALEN Expression Vectors for FLASH assembly were ordered from Addgene website. The fast ligation-based automatable solid-phase high-throughput (FLASH) assembly and expression vector construction were conducted following the protocol from Dr. Joung’s lab (Reyon et al.,

2012). 5’ end biotinylated α units were obtained first by PCR with each α unit plasmid using oJS2581 (5’-Biotin–TCTAGAGAAGACAAGAACCTGACC-3’) and oJS2582 (5’-

GGATCCGGTCTCTTAAGGCCGTGG-3’). PCR products were digested by BsaI-HF.

Extension unit and termination unit-encoding plasmids were digested with BbsI then

XbaI, BamHI-HF and SalI-HF. Digested purified extension and termination units were then ready for FLASH assembly.

Fast ligation-based automatable solid-phase high-throughput (FLASH) assembly

FLASH assembly was performed in a streptavidin-coated plate (Thermo). First, a biotinylated α unit was ligated to the first extension unit, allowing the binding of this ligated unit onto a streptavidin-coated plate in 2X B&W buffer (10 mM Tris-HCl pH 7.5,

1 mM EDTA, 2M NaCl). The following extension units and termination units were added by digesting the bead-bound fragment with BsaI-HF and an in-plate ligation with T4

DNA ligase. This process was repeated several times until the full-length of TAL fragment was obtained. Full length TAL fragment was released from the streptavidin- coated plate by digesting with BsaI-HF and BbsI. This step also created the compatible overhangs for ligating into a TALEN expression vector.

Construction of TALEN expression vectors

Expression vectors (JDS70, JDS71, JDS74, and JDS78) were digested with BsmBI and purified by AMPure XP beads (Agencourt/Beckman Genomics) according to the manufacturer’s instruction. Digested plasmids that diluted to 5 ng/μl were ready for ligation. We subcloned a full length TAL effector array into digested corresponding

TALEN expression vectors with T4 DNA ligase. Diagnostic digestion with restriction enzyme BamHI and NheI was used to determine which plasmid clone has the full-length

TALE inserted. Sequences of correct clones were confirmed by DNA sequencing using primers oSQT1 (5’-AGTAACAGCGGTAGAGGCAG-3’), oSQT3 (5’-

ATTGGGCTACGATGGACTCC-3’) and oJS2980 (5’-TTAATTCAATATATTCATGAGGCAC-3’).

Typically, in the 8 colonies we picked, there were often one or more with the correct

DNA sequence encoding 16.5 TAL effector repeats.

In vitro transcription

Expression vectors containing the ZFN or TALEN were linearized with PmeI digestion then used as template in RNA synthesis reactions using the Ambion mMACHINE Ultra-T7 kit (Forley et al., 2009). RNA was purified by lithium chloride

24 precipitation and the pellet was resuspended in 20 μl of nuclease-free water. The quality of RNAs was verified by RNA Bioanalyzer.

Microinjection

ZFN or TALEN mRNAs were mixed with phenol red and 1x Danieau solution (58 mM NaCl, 0.7 mM KCl, 0.4 mM MgSO4, 0.6 mM Ca(NO3)2, 5.0 mM HEPES pH 7.6) as previously described (Forley et al., 2009; Sander et al., 2011) (Table 4). A serial titration of RNA concentration (750 ng, 850 ng, or 900 ng per 6μl for ZFN, and 150 ng,

200 ng, or 250 ng per μl for TALEN) for each half site were tested until somatic mutation was achieved from injected embryos. For both ZFN and TALEN, we injected the solution directly into the cell at either the one-cell or two-cell stage. The volume of injection was 1 nl to 2 nl, which is about 1/10th of the total cell volume.

Genomic DNA extraction

Individual zebrafish embryos at 4 dpf were incubated in 50 μl of base solution

(25 mM NaOH, 0.2 mM Na2EDTA; pH=12) at 95°C for 30 min. After cooling down the sample, 50 μl of neutralizing solution (Tris-HCl 40 mM; pH=5) was added and mixed.

The extracted genomic DNA was spun down at 3000 rpm for 5 min every time before using for PCR (Dahlem et al., 2012).

Colony sequencing

A pool of 8 to 12 injected embryos was used for genomic DNA isolation at 4 dpf.

Embryos with abnormal appearance, for example a bent body or curled tail, were excluded from analysis. Target site regions were amplified by PCR with sequencing primers (Table 1). PCR amplicons were then subcloned into TA-TOPO PCR4

25 sequencing vectors (Invitrogen). 96 single bacterial colonies were selected and sent out for colony sequencing using the T3 priming site (5’-ATTAACCCTCACTAAAGGGA-3’) on the TOPO vector. Somatic mutation rates were calculated by percentage of colonies carrying mutations at the targeted loci.

High resolution melting analysis (HRMA)

Genomic DNAs isolated from TALEN-injected embryos or F1 embryos were examined with high resolution melting analysis according to the standard procedures provided by the manufacturer (LightCycler® 480 System) (Dahlem et al., 2012). HRMA primers specific to each target sites are listed in Table 2. Short PCR amplicons (100–

160 bp) that include the targeted region were amplified from a genomic DNA sample, subjected to denaturation and rapid renaturation. If mutations are present in the tested genomic DNA, heteroduplexes as well as different homoduplexes formed (Figure 9A).

Wild type genomic DNA, on the other hand, only contains homoduplexes after the same process. Unstable base pairing of the heteroduplexes in the mutant samples give rise to distinct melting temperatures (Tm) compared to the wild type sample (Figure 9B). LC

Green Plus dye, which fluoresces upon binding double strand DNA, was used for the detection of intact duplex molecules (Dahlem et al., 2012).

Figure 9. HRMA detects the presence of mutant alleles. (A) Schematic mechanism of HRMA. The targeted region was amplified by PCR using template genomic DNA, followed by denaturation and quick re-annealing. The genomic samples with mutant alleles produce homoduplexes as well as multiple heteroduplexes. (B) HRMA of lef1 PCR amplicons generated from either WT or heterozygous mutant. HRMA of PCR amplicons from WT template (lef1+/+, show in grey) obtained one single Tm. In contrast, re-annealed amplicons derived from the mutant (lef1Δ18/+, shown in red) consisted of multiple heteroduplexes, and thus displayed different Tms (*). Adapted from Dahlem et al., 2012. Originally published in PLoS Genetics. 8: e1002861.

Results

CoDA-ZFN induced somatic mutations in zebrafish fascin 2a gene in a site- directed manner

Zebrafish fascin 2a (EU580142.1) cDNA sequence and intron-exon junction sequences were used for selecting potential ZFN target sites via the ZiFit Targeter

(Sander et al. 2010). This software utilized its pre-screened ZFN database to provide us with several potential target sites. Each target site was designed such that the zinc- finger arrays bind specifically to 9 bp DNA on each half site. The spacer region between two half sites was 5 bp to 7 bp (Figure 10A). Target sites located at the intron-exon junction are less preferable because disruptions of splicing site are less efficient in producing a malfunctioning or truncated protein (Figure 10B). Subsequently, we used the NCBI Nucleotide BLAST tool to check the specificity of each target site in the zebrafish genome. To ensure specific targeting, sites that have possible non-specific binding that is less than 4 bp mismatch in the ZFN binding region were not used. Based on previous selection criterion, we renamed ZFN-unkown-SP-6-1 as fascin 2a ZFN6-1 for following studies. Fascin 2a ZFN6-1 target site sequence was confirmed in zebrafish genome by PCR and DNA sequencing before the construction of ZFN expression vectors.

Figure 10. CoDA target sites design for fascin 2a. (A) Potential target sites were provided by ZiFit Targeter. Colored bases are sequences of the ZF binding region. (B) Schematic representation of fascin 2a gene structure and the position of each potential target site. Introns are shown as lines, and exons are black boxes. ZFN6-1 targets the most 5’-end (to the left of the picture) of the gene. For each target half site, ZF-encoding sequence could be obtained from the ZiFit targeter (Table 3). Commercially synthesized zinc finger array-encoding DNA sequences were individually subcloned into the heterodimeric expression vectors, pMLM290 and pMLM292 (Maeder et al., 2009). Colonies with expression vectors carrying either half site were selected and confirmed by sequencing with

CMV_fwd_primer and BGH rev_primer (See Materials and Methods). In vitro transcription of ZFN-encoding RNAs was executed by using T7 RNA polymerase. The quality of the RNA produced was confirmed using Agilent Bioanalyzer virtual gels produced by the Case Genotyping Core (Figure 11). PolyA-tailed ZFN-encoding RNAs of

29 both half sites along with phenol red and 1x Danieau solution were micro-injected into one-cell staged zebrafish embryos (Table 4).

Table 3. Fascin 2a ZFN6-1 ZF-encoding DNA sequences. Sequences were obtained from ZiFit Targeter. The sequences that encode ZF1 are bold and underlined with green. ZF2-encoding sequences are bold and underlined with red. Blue sequences represent ZF3-encoding sequences. Name Sequence (5’ to 3’) Fascin 2a GAAAAAAATCTAGACCCGGGGAGCGCCCCTTCCAGTGTCGCATTTGCATGCGGAAC ZFN6-1 TTTTCGACCCGTGCAGTTTTGCGTCGTCATACCCGTACTCATACCGGTGAAAAAC Left CGTTTCAGTGTCGGATCTGTATGCGAAATTTCTCCCAGCGTTCTGACTTGACCCGT CATCTACGTACGCACACCGGCGAGAAGCCATTCCAATGCCGAATATGCATGCGCAA CTTCAGTATTCGTACCTCTTTGAAACGTCACCTAAAAACCCACCTGAGGGGATCC AAGAAGGA

Fascin 2a GAAAAAAATCTAGACCCGGGGAGCGCCCCTTCCAGTGTCGCATTTGCATGCGGAAC ZFN6-1 TTTTCGCGTCGTTTTATTTTGTCTCGTCATACCCGTACTCATACCGGTGAAAAAC Right CGTTTCAGTGTCGGATCTGTATGCGAAATTTCTCCGAAGCACATCATTTGTCTCG TCATCTACGTACGCACACCGGCGAGAAGCCATTCCAATGCCGAATATGCATGCGCA ACTTCAGTCGTTCTGACCATTTGTCTTTGCACCTAAAAACCCACCTGAGGGGATC CAAGAAGGA

Figure 10. Size of fascin 2a ZFN6-1 RNAs was confirmed by RNA Bioanalyzer. Lane 1 and 2 are fascin 2a left and right ZFN-encoding RNA (no polyA tail added), respectively. Both of them show a single band at the expected size (1033 bp). Lane 3 is polyA-tailed left ZFN-encoding RNA. Lane 4 is polyA-tailed right ZFN-encoding RNA. Size shifts were observed after polyA tailing.

Table 4. The preparation of injection solution. The amount of ZFN or TALEN-encoding RNA for each half site may vary from 750 ng to 1500 ng per 6 μl in different rounds of injections.

Component Volume Amount

RNA (Left half site) Variable 750 ng to 1500 ng RNA (Right half site) Variable 750 ng to 1200 ng 0.5% phenol red 1 μl 0.08%

1 x Danieau solution To 6 μl Genomic DNA of injected embryos was extracted on 4 dpf as a pool of 8 to 12 embryos. Injected embryos with severe developmental defects, such as a curled body, were excluded in this assay. DNA fragments containing the target site region were amplified and subcloned into sequencing vectors. Colony sequencing was used to detect mutations generated by fascin 2a ZFN6-1 RNA injection. In 96 colonies that were sequenced, one of them had an 8 bp deletion in the target site region of the fascin 2a gene (Figure 12). The amounts of ZFN-encoding RNAs injected were 1000 ng/6 μl for each half site (See Material and Methods). The somatic mutation rate of this round of injection was then determined to be about 1% (1/96), which is comparable with the somatic mutation rate in the literature (Sander et al., 2011).

Figure 12. Genotype of fascin 2a ZFN6-1 injected fish. An 8 bp deletion at the target site region was identified from the injected embryos. ZFN DNA-binding sequences are underlined.

About twenty fascin 2a ZFN6-1 injected zebrafish were raised and set up with

TÜ strain wild-type fish for identification of a founder. Individual embryos of these out- crosses were subjected to HRMA, PCR, and DNA sequencing. Unfortunately, none of the fish tested was a founder that carries mutation in the fascin 2a locus (Figure 13). This could be due to the low efficiency of ZFN-induced somatic mutation. We are now working on a more efficient gene-knockout targeting system, TALEN to generate fascin

2a knockout zebrafish.

Figure 13. No germline transmission was found for fascin 2a ZFN6-1 injected fish. This figure was generated from the HRMA of one fascin 2a ZFN6-1 injected adult zebrafish. In HRMA, amplicons derived from the genome with mutations will display distinct melting curves from the wild type control. All the genomic DNA samples from F1 embryos of this injected adult zebrafish displayed a similar melting curve to the wild type (Blue curve; thus no mutation was detected.

CoDA-ZFN induced site-specific heritable mutations in zebrafish piezo 2-4 gene

Zebrafish protein piezo 2-4, encoded by piezo 2-4 (ENSDARG00000076722), shares 75% similarity to mouse piezo 2 (NP_001034574.4). The piezo 2-4 gene has 43 exons, and the piezo 2-4 protein has 2444 amino acids and 29 predicted

32 transmembrane domains. The first transmembrane domain of the protein is encoded by part of the exon 2 sequence of the gene.

Figure 14. Location of ZFN and TALEN target sites on the piezo 2-4 gene. Above is a schematic representation of the piezo 2-4 gene structure, with the coding and the non-coding regions shown as black-filled and white-filled boxes, respectively. Introns are shown as lines. The piezo 2-4 ZFN6-1 target site sequence is in exon 4 as indicated. The piezo 2-4 TALEN1 target site is located in exon 2. The left and right ZFN or TALEN-binding DNA sequences are underlined. The procedure of selecting a piezo2-4 ZFN target site was similar to the selection of the fascin 2a ZFN target site. ZiFit Targeter (Sander et al., 2010) identified potential

CoDA ZFN target sites that bind to exons or intron-exon junctions of the gene. The binding specificity of each target site was then checked by the NCBI Nucleotide BLAST tool. Sites having potential non-specific bindings with less than 4bp mismatch were excluded from the selection. Using the above criterion, a target site in exon 4 was selected and named piezo2-4 ZFN6-1 for the following experiments (Figure 14). The target site sequence in the zebrafish genome was amplified and confirmed using our sequencing primers (Table 1).

Zinc finger array-encoding sequences were obtained from ZiFit Targeter, synthesized and ordered from a commercial resource (Table 5). DNA sequences were 33 then individually subcloned into expression vectors pMLM290 and pMLM292.

Expression vectors carrying either half site were confirmed by sequencing with

CMV_fwd_primer and and BGH rev_primer (See materials and methods). ZFN-encoding

RNAs were generated by in vitro transcription using T7 RNA polymerase. To ensure the efficiency of injection, good quality of RNAs was confirmed using the RNA Bioanalyzer

(Figure 15). Microinjection of polyA-tailed ZFN-encoding RNAs was performed on one- cell-staged zebrafish embryos (Table 4).

Table 5. ZFN-encoding DNA sequences of piezo2-4 ZFN6-1 target site. Sequences were obtained from ZiFit Targeter. The sequences that encode ZF1, ZF2 and ZF3 are bolded and underlined with green, red, and blue color, respectively. Name Sequence (5’ to 3’) Piezo 2-4 GAAAAAAATCTAGACCCGGGGAGCGCCCCTTCCAGTGTCGCATTTGCATGCGGAAC ZFN6-1 TTTTCGCAGGCATCTAACTTGACCCGTCATACCCGTACTCATACCGGTGAAAAACC Left GTTTCAGTGTCGGATCTGTATGCGAAATTTCTCCCAGCAGACCAACTTGACCCGTC ATCTACGTACGCACACCGGCGAGAAGCCATTCCAATGCCGAATATGCATGCGCAAC TTCAGTGACATGGGTAACTTGGGTCGTCACCTAAAAACCCACCTGAGGGGATCCA AGAAGGA

Piezo 2-4 GAAAAAAATCTAGACCCGGGGAGCGCCCCTTCCAGTGTCGCATTTGCATGCGGAAC ZFN6-1 TTTTCGACCCGTCAGCGTTTGCGTATTCATACCCGTACTCATACCGGTGAAAAAC Right CGTTTCAGTGTCGGATCTGTATGCGAAATTTCTCCCAGTCTGCACATTTGAAACG TCATCTACGTACGCACACCGGCGAGAAGCCATTCCAATGCCGAATATGCATGCGCA ACTTCAGTGTTCATTGGAACTTGATGCGTCACCTAAAAACCCACCTGAGGGGATC CAAGAAGGA

Figure 15. Quality of piezo2-4 ZFN6-1 RNA was confirmed by the RNA Bioanalyzer. Lane 1 is the left ZFN-encoding RNA before adding polyA tail, while lane 2 is after polyadenylation. Lane 3 is the right ZFN-encoding RNA before adding polyA tail, while lane 4 is with polyA tail added. A pool of 8 to 12 injected 4-dpf embryos, all normal looking, was used for genomic DNA isolation. The target site region was amplified and subcloned into the sequencing vector. Mutations were detected by colony sequencing as described in

Materials and Methods. Five out of 192 colonies had mutations in the target site region

(Figure 16). The somatic mutation rate was determined to be about 2.5% (5/192). For each half site, 1100 ng/6 μl ZFN-encoding RNA was injected (See Materials and

Methods).

Figure 16. Genotype of piezo2-4 ZFN6-1 RNAs injected fish. In a total of 192 colonies were sequenced, five of them had mutations in the target site region. ZFN binding sequences are underlined, while deletions are indicated by dashes. To obtain a heritable zebrafish line with the desired gene knockout, piezo 2-4

ZFN6-1 RNAs injected fish were then raised, and set up with wild-type fish for founder

(F0) screening. Genomic DNAs of individual embryos (F1) from this breeding were prepared on 4 dpf and analyzed by HRMA. Changes in the melting curve profiles

(Figure 17A, 17B) in HRMA suggested that two particular zebrafish we set up, 652.1 and 652.7 are founders that carry the mutation in the piezo 2-4 ZFN6-1 target site. To further confirm the mutation genotype, we amplified the target site region of F1 embryos by PCR and subcloned the PCR amplicons into a sequencing vector for DNA sequencing. Two F1 genomic DNAs were used in the PCR for each founder fish. For founder fish 652.1 and 652.7, five clones, out of nine we sequenced, had 4 bp deletions and 1 bp deletion in the target site region, respectively (Figure 17C, 17D). So far, we have identified two founders with successful germline transmission of the mutation in the piezo 2-4 gene locus.

Figure 17. Germline transmission of ZFN-induced mutations in piezo 2-4. HRMA successfully detected mutations in F1 embryos of fish 652.1 and fish 652.7. Amplicons derived from the genome with mutations display distinct melting curves from the wild type control. Blue curves are genomic samples exhibiting a similar melting curve to wild type, thus no mutation was detected in those samples. Melting curves obtained from embryos with mutations (other colors rather than blue) were significantly deviated from those of the WT or non-mutant fish (blue), indicating mutations at the target site. To further confirm the mutations, target site region of F1 embryos were amplified and subcloned into sequencing vectors individually. Individual clones were picked, purified and sequenced. (A) HRMA detected mutations in F1 embryos from fish 652.1, indicating that fish 652.1 is a founder. In a total of 23 F1 embryos we sent out, 12 of them had a mutation detected. (B) Germ-line transmission of fish 652.1 were confirmed by DNA sequencing. Alignment of the DNA sequencing result and predicted WT sequence revealed a 4 bp deletion at the target locus. In nine clones obtained from two F1 embryos, five of them showed the 4 bp deletion. (C) Another founder fish 652.7 identified by HRMA. In a total of 24 F1 embryos we sent out, 13 of them had a mutation detected. (D) Alignment of the DNA sequencing result and predicted WT sequence revealed a 1 bp deletion at the target locus. In nine clones obtained from two F1 embryos, five of them showed the 1 bp deletion.

TALEN induced heritable mutations in zebrafish piezo 2-4 gene with a higher efficiency

The ZFN we generated for piezo 2-4 was shown to be efficient in induction of somatic mutations as well as transmitting the mutation through germline. However, as the context-dependent property of ZFNs limits its binding site selection, the piezo 2-4

ZFN6-1 target site is on exon 4, which means that the truncated protein made from the mutation locus still has two transmembrane domains. Therefore, we took advantage of the freedom of target site selection of the TALEN system. We designed a piezo 2-4

TALEN1 target site, which is expected to disrupt more functional domain coding sequences as well as to have higher mutagenesis efficiency than ZFN.

To select a TALEN target site for piezo 2-4, exon 2 sequence of the gene was screened in ZiFit Targeter for potential sites. The criteria described in Materials and

Methods were used, and a TALEN target site was selected in exon 2 (Figure 14). This target site has been named as piezo 2-4 TALEN1 for the following studies. As this

TALEN site targets the early coding region of the gene, it is more likely to result in a piezo 2-4 null knockout than the previous ZFN we had generated.

FLASH assembly was used for the construction of TALE DNA-binding (Table

6)(Reyon et al, 2012). TALE arrays were then subcloned to respective JDS vectors as described in Table 6 and Materials and Methods. Eight colonies were picked for each half site to obtain plasmid DNA, and the correct insertion was checked by diagnostic digestion with restriction enzymes NheI and BamHI. After digestion, plasmids with the full-length TALE assembled DNA insert (~1.6kb) should have two bands at 2.4 kb and

5.6 kb, on an agarose gel (Figure 18). Sequences of positive (with full length TAL arrays) clones were then confirmed by DNA sequencing. TALEN-encoding RNAs were generated by in vitro transcription using T7 RNA polymerase and examined by the RNA

Bioanalyzer (Figure 19). PolyA-tailed TALEN-encoding RNAs were microinjected in one-cell-staged zebrafish embryos (Table 4).

Table 6. A list of TAL plasmids and JDS vectors used for the construction of piezo 2-4 TALEN. TAL plasmids were used for FLASH assembly according to the materials and methods. The full- length TALE (α unit- extension unit1 (E1)- E2- E3- Termination unit) arrays were then subcloned into respective JDS vectors. Extension Extension Extension Termination Expression α unit unit 1 unit 2 unit 3 unit vector Left TALEN TAL375 TAL182 TAL237 TAL120 TAL319 JDS74 Right TALEN TAL374 TAL37 TAL234 TAL180 TAL334 JDS71

Figure 18. Diagnostic digestion of TALEN expression vectors with full-length TAL arrays. (A) A schematic diagram of an expression vector with full length of TAL array (1.6 kb). NheI and BamHI digestion will cut out a 2.4 kb insert-containing DNA fragment from the vector (~5.6 kb). (B) An agarose gel with the digestion results of piezo 2-4 TALEN expression vectors. Positive clones having a 2.4 kb band and a 5.6 kb band after digestion are marked with arrows. Sequence of positive clones was confirmed by DNA sequencing.

Figure 19. Quality of piezo 2-4 TALEN1 RNAs were confirmed by the RNA Bioanalyzer. Lane 1 and 2 are piezo 2-4 TALEN1 left and right RNAs, respectively, before polyadenylation. Both show a single band at the expected size (3100 bp). Lane 3 and 4 are left and right TALEN- encoding RNAs after polyadenylation. Shifting of product size after polyA-tailing can be observed in lane 3 and lane 4 compared to lane 1 and lane 2, respectively. On 4 dpf, genomic DNAs of injected normal looking embryos were analyzed individually by HRMA for the presence of mutations at the target site region. In a total of 12 embryos we tested for HRMA, all of them had the melting curve shifted, and therefore had possible mutations at the target site (Figure 20A). The percentage of embryos with somatic mutations was determined to be 100% (12/12). To confirm the mutations, target regions were amplified by PCR using genomic DNAs from two injected fish, individually. PCR amplicons were subcloned into sequencing vectors. Four colonies

40 from each injected fish were purified and sequenced. From 8 plasmids, four of them from one injected fish carried, three different types of mutations (Figure 20B).

Figure 20. Induction of somatic mutation by piezo 2-4 TALEN1 in zebrafish. (A) HRMA detected mutations in TALEN-encoding RNA injected zebrafish. Genomic DNAs from individual embryos were used for HRMA. Blue curves represent those genomic samples from wild-type fish, with no mutations detected. Genomic samples with red, green and grey colors showed distinct melting curves from the control, meaning mutations were detected in those samples. In a total of 12 embryos we tested for HRM, all of them had targeted mutations detected. (B) Sequence alignment result reveals multi-forms of deletion mutations were generated in individual TALEN injected embryos at the somatic level. Around 30 injected fish were raised to adulthood. TALEN injected fish were crossed with wild type for founder (F0) screening. Genomic DNAs from individual F1 embryos were isolated at 4 dpf and subjected to HRMA. Changes of melting curve of one embryo suggested that 701.3 is a F0 founder fish that carries mutations at the targeted locus (Figure 21A). We then amplified the target site region of two F1 embryos individually and subcloned the PCR amplicons into sequencing vectors. Four colonies were sequenced for each F1 embryo. All four plasmids from one embryo had a 3 bp insertion in the piezo 2-4 TALEN targeted region, which confirmed that 701.3 is a founder fish (Figure 21B). In a total of five adult fish we have screened so far, one founder fish with successful germ-line transmission was found.

Figure 21. Germline transmission of TALEN-induced mutations in piezo 2-4 gene was found in fish 701.3. (A) Mutations in F1 embryos were detected by HRMA. Blue curves are genomic samples exhibiting a similar melting curve to the wild type, thus no mutation was detected in those samples. Red curve is a genomic DNA sample with mutation detected. In a total of 8 F1 embryos we analyzed, one of them had mutation detected. (B) Alignment of the DNA sequencing result revealed a 3 bp insertion at the target locus. All four colonies from one F1 embryos showed the same 3 bp insertion. TALEN binding region were underlined, and insertions were highlighted in red.

Discussion

Zebrafish reverse genetics has lagged behind that of other model systems. Now it seems possible to bring zebrafish genetics to the forefront. As our goal is to build a platform for generating knockout zebrafish by ZFN and TALEN to study molecular mechanisms of hair-cell development and mechanotransduction, we have designed and generated ZFNs and TALENs for 18 hair cell expressed genes. In summary, we have successfully set up a platform for generating knockout zebrafish using ZFN and TALEN technologies (Figure 22). We showed that somatic mutation rates obtained in this study are comparable to the published data (Dahlem et al., 2012; Sander et al., 2011).

The somatic mutation rates of ZFN injected embryos ranged from 0.5% to 3% as shown in results (detected by colony sequencing). As our skill developed, HRMA became a simpler and cost-effective method for us to detect mutation (Dahlem et al., 2012).

However, our previous results suggested that CoDA ZFN induced somatic mutation rates are too low to be detected by HRMA if not using a large number of fish for examination. The percentages of TALEN injected embryos having somatic mutation detected by HRMA are above 50%, which is also comparable with data from other research groups (Moore et al., 2012; Cade et al., 2012). Moreover, several founder fish that successfully transmit the mutations at the target site to its offspring are found in our ZFNs or TALENs injected fish. The germ-line transmission rates are around 6.7%

(2/30) for ZFNs injected fish, while 20.0%(1/5) for TALENs injected fish. We therefore concluded that TALEN has a higher success rate than ZFN in inducing somatic mutations as well as germline transmission in zebrafish.

Figure 22. Flow chart of generating knockout zebrafish using ZFN or TALEN. Time needed for each step is shown in red. By our ZFN or TALEN platform, somatically mutated zebrafish can be generated in two months. A founder fish may be obtained in 6 months. To predict how the mutations we obtained would affect Piezo 2-4 protein function, we did translation analysis on the cDNA sequences after deletions or insertions at the targeted loci using Macvector Software. Mutations in piezo 2-4 ZFN6-1 founder fish 652.1 and 652.7 give rise to early stop codons, further results in generation of truncated Piezo 2-4 proteins with 181 amino acids and 182 amino acids long respectively. The full length of Piezo 2-4 protein is 2444 amino acids. For piezo 2-4

TALEN induced mutations, the 3 bp insertion shown in Figure 21 is an in-frame insertion with the addition of one amino acid to the wild-type protein. However, previous published data demonstrated that a single founder fish generated by TALEN might carry different mutant alleles (Huang et al., 2011; Cade et al., 2012). Thus,

44 founder fish 701.3 should be considered as a mosaic, which has multi-mutant genotypes and it is highly possible to carry other types of heritable mutations in piezo 2-4 TALEN target site that will produce truncated Piezo 2-4 protein.

As one of the first technologies available to induce mutations in specific zebrafish genes, ZFN greatly facilitated the projects on genomic manipulation in zebrafish field. I have demonstrated that CoDA ZFNs are effective in inducing heritable mutations at the selected targeted loci. However, the pre-selected zinc finger units that each bind to 3 bp DNA sequences largely limits the targeting range of CoDA ZFN for a given gene. In contrast, TALEN can be designed to target almost any site. Construction of TALENs can be done by a rapid and low-cost method, FLASH assembly as described in the Material and Method (Reyon, et al., 2012). Thus even though both platforms were shown to be effective in our study, TALENs are more likely to be widely used for the site-directed zebrafish mutagenesis than CoDA ZFNs.

Another system that can be used to knockout specific genes in zebrafish is called the CRISPR-Cas system (Hwang et al., 2013). In this system, customizable single guide

RNAs (sgRNAs) direct the Cas9 endonuclease to the target locus and mediated gene- specific alteration. CRISPR-Cas system not only can target endogenous genes in zebrafish, but also were shown to be efficient in simultaneous disruption of multiple genes in mouse embryonic stem cells (Yang et al., 2013). The mutation frequencies induced by CRISPR-Cas system in targeted zebrafish genes were reported to be from

27.1% to 59.4%, which is comparable to TALEN (Hwang et al., 2013). The productions of sgRNA and Cas9 RNA are simpler than construction of ZFN or TALEN. However, this system now can only target sequences with the format of 5’-(G/A)(G/A)-N18-NGG-3’,

45 which occurs once in every 32 bp of random DNA sequences. In addition, recent published data from Fu and colleges demonstrated that CRISPR-associated Cas9 based

RNA guided nucleases targeted endogenous loci in human cells with a high off-targeting mutagenesis rate (Fu et al., 2013). As a relatively new gene-targeting technology, the off-targeting effects of this system in zebrafish still need to be determined. Therefore, I am currently working on targeting one or multi-gene loci in zebrafish using CRISPR-

Cas9 system.

Together, these genome editing techniques, ZFN, TALEN, and CRISPR enable the generation of gene-specific knockout zebrafish in the laboratory and allow us to decipher the complex molecular mechanisms of hair bundle genesis and mechanotransduction. Upon obtaining the knockout zebrafish line for our targeted genes, the function of a protein in hair-cell morphologenesis could be studied by immunostaining with a combination of antibodies. In addition, several functional assays can also be performed in these homozygous knockout zebrafish, such as (1) acoustic startle response (Bang et al., 2002), (2) cell labeling by FM1-43 dye (Gale et al., 2001),

(3) imaging of genetically encoded GCaMP calcium indicator (Nakai et al., 2001), (4) patch clamp techniques for channel studies in hair cells (Einarsson et al., 2012). The first three assays are used to test the mechanotransduction function of hair cells. Patch clamp experiments, on the other hand, will allow us to resolve the details of defects in channel properties in gene-specific knockout hair cells.

As no founder fish were obtained from fascin 2a ZFN injected fish, we are working on knocking out fascin 2a by TALEN technology. I expect that the knockout zebrafish will have deformed hair bundles: thin, short or fused stereocilia. Due to the

46 impairment of hair bundles, decreased acoustic startle responses might be observed.

For piezo 2-4 gene, the next step will be to cross F1 heterozygous with each others and identify any phenotypical changes related to changes of genotypes in their offspring using assays mentioned above. Meanwhile, offspring of the cross should be raised and screened for homozygous mutant F2 for future studies. In our hypothesis, if Piezo 2-4 protein is the mechanotransduction channel or a member of the mechanotransduction apparatus, knockout zebrafish with truncated Piezo 2-4 proteins should interrupt the process of mechanotransduction in hair cells. Therefore, we would expect a weaker or no response to acoustic startle response testing, no FM1-43 dye loading, and no influx of calcium as an indication of the impairment of the functional mechanotransduction apparatus in piezo 2-4 knockout zebrafish.

In summary, this thesis shows that I am able to set up ZFNs and TALEN based gene-knockout platforms to knockout specific genes in zebrafish. By targeting hair cell- related genes in zebrafish, the platforms I developed will facilitate our studies in understanding the molecular mechanisms of zebrafish hair cell genesis and functions.

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