PROTOCADHERIN-17 FUNCTION
IN ZEBRAFISH RETINA DEVELOPMENT
A Dissertation
Presented to
The Graduate Faculty of The University of Akron
In Partial Fulfillment
of the Requirements for the Degree
Doctor of Philosophy
Yun Chen
December, 2012
PROTOCADHERIN-17 FUNCTION
IN ZEBRAFISH RETINA DEVELOPMENT
Yun Chen
Dissertation
Approved: Accepted:
______Advisor Department Chair Dr. Qin Liu Dr. Monte Turner
______Committee Member Dean of the College Dr. Richard Londraville Dr. Chand Midha
______Committee Member Dean of the Graduate School Dr. Brian Bagatto Dr. George R. Newkome
______Committee Member Date Dr. Bruce Cushing
______Committee Member Dr. Zhonghui Duan
ii
ABSTRACT
Expression and function of protocadherin-17 (pcdh17) in zebrafish retinal development were analyzed in this study. Pcdh17 mRNA (pcdh17) expression pattern was characterized using whole mount in situ hybridization method. Antisense morphlino oligonucleotides (MOs) technique was used to determine pcdh17 function. Moreover, molecular mechanisms underlying pcdh17 expression and function were studied using proteomics. pcdh17 was expressed in developing zebrafish retina during critical stages of its development. Abnormal eye and retinal development was observed in developing zebrafish (49-72 hours post fertilization, hpf) injected with zebrafish pcdh17 specific
MOs (pcdh17 morphants). The morphants had significantly smaller eyes and disrupted differentiated retinal cells (e.g. retinal ganglion cells and photoreceptors), due mainly to decreased cell proliferation as well as defects of nonneuronal cell differentiation.
Proteomic analysis revealed that several hundred proteins were differentially expressed between wildtype embryos and pcdh17 morphants. A subset of spots showing the biggest differences were identified using Mass Spectrometry. These proteins included phosphoglycerate kinase, beta-actin-like protein, and glial fibrillary acidic protein
(GFAP). Most of the identified proteins are involved in basic cellular metabolism and cellular structure, whereas GFAP is a molecule involved in Notch signaling pathway known to play a critical role in vertebrate retinal development. In addition, pcdh17 iii
functions in retina development via regulating certain transcription factors, classical cadherins, homophilic reacting molecules and integrin-dependent adhesion. My research suggests that pcdh17 plays an important role in zebrafish retinal development, likely through involving multiple pathways including Notch-Delta, Wnt pathways.
iv
ACKNOWLEDGEMENTS
I would never have been able to finish my dissertation without the guidance of my committee members, help from friends, and support from my family.
I would like to express my deepest gratitude to my advisor, Dr. Qin Liu, for his excellent guidance, caring, patience, and providing me with an excellent atmosphere for doing research. I would like to thank Dr. Richard Londraville, who taught me the proteomics in past year. He also helped me to improve my scientific writing from the first year I was in US. It is a great technical support for my research. I would also like to thank
Dr. Brian Bagatto and Dr. Bruce Cushing and Dr. Zhong-Hui Duan for guiding my research helping me to develop my background in physiology, neuroscience and computational biology. Also a thank to Dr. Jun Hu with his help in chemistry field.
My colleagues as well as friends, Mark Dalman, Hope Ball, Donald Copeland,
Sunil Bhattarai and Alicja Sochacka were always willing to help and give their best suggestions. My research would not have been possible without their help.
I would also like to thank my family. They were always supporting me and encouraging me with their best wishes. They were always there cheering me up and stood by me through the good times and bad.
My research is supported by grants from National Institutes of Health (R15
EY013879). v TABLE OF CONTENTS Page
LIST OF TABLES ...... vii
LIST OF FIGURES ...... viii
CHAPTER
I. INTRODUCTION ...... 1
II. LITERATURE REVIEW ...... 4
III. PCDH17 mRNA EXPRESSION IN DEVELOPING ZEBRAFISH RETINA ...... 13 Materials and methods ...... 13 Results ...... 16
IV. STUDY OF PCDH17 FUNCTION IN ZEBRAFISH RETINAL DEVELOPMENT 18 Materials and methods ...... 18 Results ...... 24
V. MOLECULAR MECHANISM UNDERLYING pcdh17 FUNCTION ...... 39 Materials and methods ...... 39 Results ...... 43
VI. DISCUSSION AND CONCLUSIONS ...... 57 Pcdh17 is essential in zebrafish eye development ...... 57 Loss-of-function techniques ...... 60 Possible mechanisms of pcdh17 function in retinal development ...... 62 Summary and perspective ...... 66
REFERENCES ...... 68
vi LIST OF TABLES
Table Page
1. Effects of pcdh17 MOs injections on zebrafish development ...... 28
2. Effects of pcdh17 MOs injections on eye size ...... 29
3. Effects of pcdh17 knockdown on zebrafish retinal development ...... 32
4. Effects of vivo-pcdh17sMO injection on eye size ...... 36
5. Effects of vivo-pcdh17sMO injection on zebrafish retinal development ...... 38
6. Effects of pcdh17 knockdown on expression of transcription factors ...... 45
7. Identification parameters for zebrafish MS analyzed protein bands ...... 49
8. Gene ontology Biological Process Terms of the MS identified proteins ...... 53
9. Gene ontology Molecular Function Terms of the MS identified proteins ...... 54
10. Gene ontology Cellular Component Terms of the MS identified proteins ...... 55
vii LIST OF FIGURES
Figure Page
1. The dual function of β-catenin in cell adhesion and transcription ...... 6
2. Expression of pcdhs from genomic DNA to functional protein ...... 8
3.Wide-type 72 hpf zebrafish eye...... 9
4. pcdh17 expression in the developing zebrafish retina ...... 17
5. Gross morphological defects in pcdh17 morphants ...... 25
6. Higher magnification of eye regions of embryos ...... 25
7. Diagnostic RT-PCR demonstrating efficacy of the splice-blocking pcdh17 MO...... 26
8. Immunostaining analysis of retinal cell differentiation ...... 30
9. Retinal organization revealed by beta-catenin immunostaining...... 31
10. Apoptosis analysis using TUNEL staining ...... 33
11. Histone-H3 immunostaining ...... 34
12. Gross body and eye morphology in live control and vivo-morphant embryos...... 35
13. Immunostaining analysis of retinal cell differentiation in vivo-morphant retina ...... 37
14. Expression of transcription factors ...... 46
15. Blue stained gels of total proteins of developing zebrafish ...... 47
16. Coomassie blue staining 2-DE electrophoresis map...... 48
17. Immunohistochemstry confirmation of GFAP expression ...... 56
viii CHAPTER I
INTRODUCTION
Cadherins are cell adhesion molecules that play crucial roles in vertebrate development (Takeichi, 1990). During development of multicellular organisms, cadherins are involved in both cell-cell adhesion and cell signal transduction (Takeichi, 1988;
Huelsken and Birchmeier, 2001; Yamaguchi, 2001). All cadherins share a similar structure: a large extracellular (EC) domain, a transmembrane domain and a cytoplasmic domain (Suzuki, 1996). According to differences in these domains and genomic organization, more than one hundred cadherins have been divided into several subfamilies including classical cadherins, desmogleins, desmocollins and protocadherins
(Suzuki, 1996; Nollet et al., 2000).
Classical cadherins are well known in cell-cell adhesion, cytoskeleton anchoring and signal transduction (Takeichi, 1991, 1995; Gumbiner, 2000; Yagi and Takeichi,
2000). Numerous classical cadherins are involved in development of tissues and organs and maintenance of adult structures (Takeichi, 1995; Gumbiner, 2000). E-cadherin is mainly expressed in epithelial tissues and cell adhesion mediated by homophilic interactions between E-cadherin expressing cells plays an essential role in early embryonic development: disruption of E-cadherin function results in disintegration of embryos at blastula and gastrula stages (Hirai et al., 1989; Wheelock and Jensen, 1992; 1 Perrais et al., 2007; Syed et al., 2008). E-cadherin function is also required for adult animals, and reduced function has been implicated in tumor formation and pathogenesis of many forms of cancers in both animal models and humans (reviewed by Wijnhoven et al., 2000). N-cadherin is mainly expressed on the cells of developing central nervous system (CNS) and blocking its expression results in massive morphological changes in the CNS (Hatta and Takeichi, 1986; Ganzler and Redies, 1998). Similar CNS defects are also found in zebrafish N-cadherin mutants (Lele et al., 2002). Differentiation of several major retinal cell types including retinal ganglion cells (RGCs) also requires involvement of classical cadherins (Malicki et al., 2003; Masai et al., 2003; Babb et al., 2005; Ruan et al., 2006; Liu et al., 2007; Liu et al., 2008). N-cadherin is crucial in early retinal neuroepithelium development, and later in differentiation of RGCs and photoreceptors
(Malicki et al., 2003; Masai et al., 2003; Liu et al., 2007). R-cadherin (cdh4) is critical for RGCs differentiation (Babb et al., 2005). Knockdown of K-cadherin (cdh6) caused retinal defects in both zebrafish and frogs (e.g. smaller eyes, disrupted RGCs and amacrine cell differentiation), indicating that cdh6 plays a key role in normal formation of the retina (Ruan et al., 2006; Liu et al., 2008).
Protocadherins (pcdhs) are the biggest subgroup containing more than 80 members. Some pcdhs were found to be expressed in the developing retina. Pcdh1, one of the first pcdhs discovered, was shown to be expressed in the presumptive ganglion cell layer and inner nuclear layer of mouse retina during its embryonic development (Redies et al., 2008). Pcdh9 was not detected in early zebrafish embryonic retina but in older ones
(50 hpf-72 hpf, Liu et al. 2009). At 50 hpf, it was seen in the retinal ganglion cell layer in
2 the ventroanterior region of the retina where differentiation of retinal cells is initiated (Hu and Easter, 1999). At 72 hpf, its expression had expanded to the entire retinal ganglion cell layer (Liu et al. 2009). Pcdh18 was observed in the developing frog’s retina (Kubota et al., 2008), while pcdh19 was detected in developing zebrafish retina (Liu et al., 2010).
Despite knowledge of expression of several pcdhs in the developing vertebrate retinas, there is little known about pcdhs functions the retinal development. In this study,
I focused on the study of protocadherin-17 (pcdh17, also called protocadherin-68) function in zebrafish retinal development. Unlike many classical cadherins and some pcdhs which are expressed more widely (e.g. the entire organism or the nervous tissue), pcdh17 expression was more limited (e.g. in developing retina and parts of the brain). It has been isolated from several species, including humans, rat and zebrafish (Kim et al.,
2007; Biswas and Jontes, 2009; Liu et al., 2009). Using zebrafish as the model organism,
I attempt to determine pcdh17 expression in the developing zebrafish retina, study pcdh17 function zebrafish retinal development and gain insights of molecular mechanism underlying pcdh17 function in the retinal development.
3 CHAPTER II
LITERATURE REVIEW
All cadherins have a large extracellular (EC) domain and a transmembrane domain, and most cadherins also contain a cytoplasmic domain. In classical cadherins, the EC domain consists of five homologous repeats of a cadherin motif, with each repeat showing characteristic features in addition to some common properties. For example, classical cadherins mainly bind to other cadherin molecules of the same type on adjacent cells (homophilic interactions). This type of interaction is based on two functional structures in their EC repeat 1 (EC1), tryptophan 2 (W2) and a hydrophobic pocket.
Hydrophobic pocket insert in adjacent cadherin W2 site to form homophilic cell-cell adhesion (Boggon et al., 2002).
Transmembrane domain anchors the cadherin molecule to the cell membrane.
The cytoplasmic domain is the most conserved part of the cadherin molecule, suggesting the functional importance of this region. For classical cadherins, this domain plays a role in cytoskeleton anchoring and signal transduction by reacting with a complex of proteins
(e.g. β-catenin), which in turn can become associated with the actin microfilament system within the cell. Cadherin-β-catenin-actin interaction can affect cell structure, while cadherin-β-catenin-other proteins (e.g. T-cell factor transcription factor) interactions affect cell differentiation and proliferation (Takeichi, 1991; Brembeck et al.,
4 2006). Figure 1 shows a trigger role of E-cadherin in canonical Wnt pathway. E-cadherin combines with β-catenin, which further links to α-catenin. If β-catenin is phosphorylated by tyrosine kinases Fer, Fyn or Met, it can lead to a loss of E-cadherin-binding and the
“released” β-catenin can interact with the nuclear co-factor BCL9-2. The β-catenin-
BCL9-2 complex regulates, in conjunction with the LEF/TCF DNA binding proteins, transcription of crucial target genes (Clevers, 2006). If β-catenin is degraded by a destruction complex including adenoma polyposis coli (APC) gene product, scaffold molecules axin (conductin ortholog, also known as axin2), glycogen synthase kinase
(GSK3β) and casein kinase (CKI, Brembeck et al., 2006). Thus, the adhesive ability and movement of cells as well as gene transcription are inhibited.
5
Figure 1. The dual function of β-catenin in cell adhesion and transcription (Brembeck et al., 2006).
Unlike classical cadherins, pcdhs have six or seven EC repeats instead of five; having neither W2 nor hydrophobic pocket, but having a characteristic disulfide-bonded loop in the EC1 domain (Morishita and Yagi, 2007). The disulfide-bonded loop is located close to the clusters of calcium-binding residues near the N terminus of pcdhs and may participate in interactions with other proteins, such as with glycoproteins rather than adjacent cadherins (Perez and Nelson, 2004; Morishita et al., 2006). Pcdhs also have a
RGD motif in the EC domain (EC1 or EC2) which is essential for integrin-dependent cell adhesion (heterophilic adhesion activity, Kim et al., 2011). The cytoplasmic region of pcdhs does not have catenin binding sites, suggesting that pcdhs may mediate cell-cell
6 adhesion and/or regulating cell signal transduction by interacting with other cytoplasmic proteins such as phosphorylated serine/threonine phosphatase type 1 (Fyn, Kohmura et al., 1998; Yoshida et al., 1999) and phosphotyrosine-binding domain of the adaptor protein Disabled 1 (Homayouni et al., 2001).
Several subgroups of pcdhs have been identified, including α-, β-, γ- and δ-pcdhs.
The first three are encoded by three gene clusters on chromosome 5q31 in humans (Wu et al., 2001; Frank and Kemler, 2002). These pcdhs are called clustered pcdhs. In contrast,
δ-pcdhs are non-clustered, and encoded by genes located on different chromosomes
(Wolverton and Lalande, 2001; Redies et al., 2005; Vanhalst et al., 2005). The δ-pcdhs can be further divided into δ1-pcdhs (e.g. pcdh1, 7, 9) and δ2-pcdhs (e.g. pcdh17, 18, 19), based on the number of EC repeats and conserved motifs (CMs) in the cytoplasmic domain. The δ1-pcdhs have seven EC repeats and three CMs (CM1-CM3), whereas δ2- pcdhs contain six EC repeats and only two CMs (CM1 and CM2, Redies et al., 2005).
In classical cadherins, multiple exons are corresponding for whole protein encoding. By contrast, all pcdhs share a similar genomic structure that a single large exon
(exon 1) encodes for the entire extracellular domain, as well as the transmembrane domain and a small part of the cytoplasmic domain (Wu and Maniatis, 2000, Fig. 2).
7
Figure 2. Expression of pcdhs from genomic DNA to functional protein. Abbreviations: EC, extracellular domain; TM, transmembrane domain; CP, cytoplasmic domain.
Pcdh17 is a member of the δ2-pcdhs and mainly expressed in developing CNS including the retina (Biswas and Jontes, 2009; Liu et al., 2009). There are several reports about its function in tumor-suppressive activity in humans (Haruki et al., 2010; Hu et al.,
2012). However, there is little information on its function in the vertebrate development and molecular adhesion activity.
Zebrafish (Danio rerio) was used as the model organism. This model was first utilized by Streisinger and his colleagues at the University of Oregon to study mutants among parthenogenetic offspring of mutagenized females (Streisinger et al., 1981). The fish is easily raised and maintained, kept in large numbers and used in embryological manipulations. Its genomic information is available in online databases (such as ZFIN and NCBI). Studies have convincingly demonstrated that molecular mechanisms underlying animal development are remarkably similar (Alberts et al., 2002). In addition 8 anatomy and physiology of zebrafish are well characterized and similar to other vertebrate animals, e.g. mice and humans (Cepko, et al., 1996; Schmitt and Dowling,
1999). Thus, by understanding how genes function in zebrafish, we may be able to gain insights about their roles in humans.
Zebrafish retina develops in a inside first, outside last fasion, forming several distinct layers at 72 hours post fertilization (hpf, Fig. 3, Avanesov and Malicki, 2004).
Retinal ganglion cells (RGCs) begin to differentiate around 26-28 hpf in the anteroventral part of the retina adjacent to the lens (Nawrocki, 1985). Axons of differentiating RGCs exit from the retina as an optic nerve, arriving at the optic chiasm around 32 hpf, and cover its main brain target, the optic tectum, by 72 hpf (Nawrocki,
1985; Stuermer, 1988; Burrill and Easter, 1994).
Figure 3. Wide-type 72 hpf zebrafish eye. This is a cross section with dorsal to the top (Avanesov and Malicki, 2004). Abbreviations: gcl, ganglion cell layer; inl, inner nuclear layer; ipl, inner plexiform layer; mz, marginal zone; on, optic nerve; opl, outer plexiform layer; pcl, photoreceptor layer; pe, pigmented epithelium.
9
Amacrine cells, the second type of retinal cells, start to develop around 40 hpf
(Schimitt and Dowling, 1999). The neuronal layer where amacrine cells reside is called inner nuclear layer (inl). As development continues, a plexus of neurites is formed projecting toward the gcl (Godinho et al., 2005). By 50 hpf, dendrites of RGCs and amacrine cells form a dense reticulum of fibrils, called inner plexiform layer (ipl).
In addition to the amacrine cells in the innermost part, the inl also contains horizontal cells (outer portion) and bipolar cells (between the other two). Horizontal cells begin to differentiate around 57 hpf, and bipolar cells begin to differentiate at about 60 hpf (Hu and Easter, 1999; Li et al., 2000).
Photoreceptors are located in outermost potion of the retina. The earliest photoreceptors begin to form around 45 hpf (Raymond et al., 1995; Bilotta and Saszik,
2001). By 72 hpf, they expand to the entire retina and form a distinct layer called photoreceptor layer (pcl, Malicki, 1999). During this time, dendrites of horizontal cells and bipolar cells from the inl, and photoreceptors from pcl project to each other to form a thin fibrous layer called outer plexiform layer (opl), which is a synaptic layer located between the inl and pcl (Masland, 2001; Moorman, 2001).
Besides those types of retinal neurons, nonneuronal glial cells (e.g. Müller glial cells) also play important roles in the retina development. A previous study showed that this type of cells was involved in initial patterning of axonal pathways in the CNS
(Marcus and Easter, 1995). Radial glia of the retina support neurons, provide nutrition to retinal neurons and are involved in retinal cell differentiation (Bringmann et al., 2006).
10 Moreover, Müller glia cells may also function as retinal stem cells and guide migration of progenitors into the retinal neuron layers (Raymond et al., 2006).
To explore pcdh17 function in development of these retinal cells, pcdh17 gene was knocked down using antisense morpholino oligonucleotide (MO) techniques, which have been used to block gene function in vertebrates effectively and selectively in several animal organisms, such as Xenopus and zebrafish (Ekker, 2000; Nasevicius and Ekker,
2000).
Han et al. (2010) used proteomics to profile γ-pcdh (a clustered pcdh) protein complexes, showing that pcdhs might function in multiple intracellular signaling pathways. Proteomics were also performed in this dissertation to study the molecular mechanisms underlying pcdh17. Proteomics simply can be defined as an efficient study of a genome’s protein expression (Blackstock and Weir, 1999). It characterizes the entire protein complement of a cell, tissue or organism to obtain a global and integrated view of expression on a large scale (Wilkins et al., 1996). Proteomics was developed to assay protein expression, protein-protein interactions, posttranslational modifications of proteins and alteration of protein composition in tissues, which can provide more informative insights into gene functions, unlike genomics-based approaches (Banks et al.,
2000). Two key tools are used typically in proteomics: 1) Two-dimensional sodium dodecyl sulfate polyacrylamide gel electrophoresis (2D SDS-PAGE, O’Farrell, 1975) and
2) Mass spectrometry (MS). The 2D SDS-PAGE allows a routine separation of 5,000 to
10,000 protein spots per gel for analysis. Modern MS techniques were a major breakthrough for protein identification. It is highly accurate and efficient and has
11 essentially replaced classical Edman sequencing as the protein identification tool of choice (Andersen and Mann, 2000).
Proteomics was first used by Wilkins et al. (1996) to profile protein complement of E. coli. It since has been widely used in clinical and biomedical applications including drug development, assessments of neurological disorders, infectious diseases, heart disease and cancer (Banks et al., 2000; Dunn, 2000; Wittmann-Liebold et al, 2006). The
2D SDS-PAGE and MS techniques have also been used to document protein expression profiles in developing zebrafish (10, 48, 72 and 120 hpf separately, Link et al., 2006;
Lucitt et al., 2008). Moreover, the developing zebrafish protein expression profiles change in response to changes in environment and/or levels of toxic chemicals (Shi et al.,
2009; Hanisch et al., 2010).
With rapid advancements in MS and bioinformatics, proteomics has become a powerful tool for analyzing signaling pathways. I therefore used proteomics to probe molecular mechanisms underlying pcdh17 function in zebrafish retinal development.
12 CHAPTER III
PCDH17 mRNA EXPRESSION IN DEVELOPING ZEBRAFISH RETINA
Pcdh17 mRNA (pcdh17) was found to be expressed in embryonic rat retina (Kim et al., 2007). Knowledge of spatial and temporal expression patterns of pcdh17 in the embryonic zebrafish retina would be a necessary first step in determining pcdh17 function in zebrafish retinal development.
Materials and methods
Zebrafish (Danio rerio). Zebrafish embryos were obtained by breeding in house wildtype adult zebrafish. The embryos were placed in filtered fish tank water in 400 ml plastic beakers kept at 28.5oC, 12 hour light and 12 hour dark. Embryos used for whole mount in situ hybridization (WISH) or whole mount immunocytochemistry (WICC) were raised in phenylthiourea (PTU, 0.003%) treated fish tank water to inhibit pigmentation.
The embryos were allowed to develop to desired stages (e.g. 24 hpf), killed using an overdose of anesthetizing agent MS-222, and fixed in 4% paraformaldehyde in phosphate buffered saline (PBS, pH=7.4) overnight at 4oC. The next morning, embryos were washed in PBS three times, 10 min. each. The embryos were incubated with increasing concentrations of methanol and finally placed in 100% methanol and stored at -20oC until use. Detailed procedures for zebrafish care and tissue processing were described in the 13 Zebrafish Book (Westerfield, 2005). All animal related protocols have been approved by the Animal Use and Care Committee at the University of Akron.
Whole mount in situ hybridization (WISH). Total RNA was isolated from 45-72 hpf zebrafish embryos using Trizol reagent (Invitrogen, Carlsbad, CA), and reverse transcriptase-polymerase chain reaction (RT-PCR) was performed using zebrafish pcdh17 specific primers (forward primer 5'-ctgtgtttgaacagccctca-3'; reverse primer 5'- ttgcaccatcagtgggttta-3') to obtain a pcdh17 complementary DNA fragment (cDNA, corresponding to the nucleotides 678–1525 of the zebrafish pcdh17, GenBank Accession
No. XM_684743). Resultant cDNA, cloned into a pCRII-TOPO vector (Invitrogen,
Carlsbad, CA), was verified by restriction mapping and a PCR experiment using a pair of pcdh17 specific primers that were internal to the set of the above primers (forward primer
5'-gaagggattaatgggcaggt-3'; reverse primer 5'-ccagaacagaacccaggtattc-3'). The confirmed pcdh17 cDNA fragment (approximately 1 μg) was transcribed with T3 polymerase to generate antisense cRNA probes using the DIG RNA Labeling Kit (Roche, Indianapolis,
IN). Procedure for synthesis of digoxigenin (Dig)-labeled cRNA probes was previously described (Liu et al., 1999). Detailed procedures for WISH were reported in Liu et al.
(1999). Briefly, fixed embryos were rehydrated in decreasing concentrations of methanol in PBST (PBS supplemented with 0.1% Tween 20). The embryos were treated with proteinase K (Roche, 10 ug/ml in PBST), re-fixed in 4% paraformaldehyde, washed in
PBST, and placed in a hybridization solution (1% 1.0 M Tris, pH7.5; 50% formamide; 6%
5.0 M NaCl; 0.2% 0.5 M EDTA; 10% Dextran Sulfate; 1% Blocking solution). The embryos were incubated with the hybridization solution at 58oC for 2 hours, followed by
14 replacing the hybridization solution with a fresh hybridization solution containing the
Dig-labeled pcdh17 cRNA probes (about 1 ug/ml). The embryos were hybridized overnight at 58°C. The next morning, embryos were washed in a series of solutions
(2xSSC, fermamide/2xSSC, 0.2xSSC, PBST), followed by incubating in a blocking solution (1% normal goat serum, 0.2 g/l ml Bovine Serum Albumin Fraction V (BSA,
Roche), and 1% DMSO) at room temperature for 2 hours. The blocking solution was replaced by a fresh blocking solution containing an alkaline phosphatase-conjugated Fab anti-DIG fragments (1:5,000, Roche), and the embryos were incubated in the solution overnight at 4°C on a shaker. For visualization of labeled mRNA, the embryos were washed in PBST, followed by genius buffer 3 (Roche, pH = 9.5), and treated with a solution containing 5-bromo-4-chloro-3-indoyl phosphate (BCIP) and nitroblue tetrazolium salt (NBT, one NBT/BCIP tablet (Roche) dissolved in 10 ml distilled water) in dark until satisfactory color reaction was achieved (about 1 hour). The embryos were then washed in PBST, fixed again in 4% paraformaldehyde for 20 min. at room temperature. After AN additional two washes in PBST, the embryos were subsequently placed in 50% glycerol in PBS and stored at -20oC till used for microscopy.
Microscopy and photography. Processed embryos were viewed under an Olympus compound microscope (BX51, Olympus America Inc., Center Valley, PA) equipped with a SPOT digital camera (SPOT imaging Solutions, Sterling Heights, MI). Digital images were taken using the fluorescent or Nomarski optics. Digitized images were processed with the same values of contrast, sharpeness, and hue/saturation with Adobe Photoshop
(Adobe Systems, San Jose, CA).
15 Results
Using WISH with pcdh17 cRNA probes, pcdh17 expression in developing zebrafish retina was studied (Fig. 4). At 26 hpf when pcdh17 was first detected in the embryonic retina, pcdh17 was found in a small region located in the anteroventral retina
(Fig. 4A). By 34 hpf, it was expressed in the entire retina (Fig. 4B) as retinal cells in the inner most region (adjacent to the lens) begin to differentiate into retinal ganglion cells
(RGCs, Nawrocki, 1985; Hu and Easter, 1999). By 50 hpf, differentiated RGCs occupy a distinct layer adjacent to the lens, retinal cells in the inner portion of the inner nuclear layer (inl) begin to differentiate into amacrine cells, and some photoreceptors have begun to form in the outer nuclear layer (onl, Robinson et al., 1995; Schmitt and Dowling,
1996, Hu and Easter, 1999; Godinho et al., 2005). pcdh17 expression pattern in the retina remained similar to that at 34 hpf (Fig. 4C), except that cells in the outer portion of the inl showed stronger pcdh17 expression. All retinal layers are well formed in 72 hpf embryos. At this stage, pcdh17 expression was still detected throughout the retina, but the outer portion of the inl, where horizontal cells reside, showed the highest level of pcdh17 expression, while its expression in the inner regions of the retina was reduced
(Fig. 4D). In addition, the newly developed onl where differentiating photoreceptors are located had the reduced pcdh17 expression (Fig. 4D).
16
Figure 4. pcdh17 expression in the developing zebrafish retina. All panels show lateral views of whole-mount retinae (anterior to the left and dorsal up) processed for pcdh17 in situ hybridization. The arrow in panels A and B indicates the retinal pigmented epithelium. The arrowhead in panel pcdh17 expression in the developing zebrafish retina A points to pcdh17 expression in the anteroventral retina. Abbreviations: gcl, ganglion cell layer; inl, inner nuclear layer; le, lens; nr, neural retina; onl, outer nuclear layer. Scale bars = 50 μm for all panels.
17 CHAPTER IV
STUDY OF PCDH17 FUNCTION IN ZEBRAFISH RETINAL DEVELOPMENT
Two types of antisense morpholino oligonucleotides (MOs) were used to knockdown pcdh17 function, followed by analyzing eye size, morphology, and differentiation of selective retinal cells using retinal/neural markers. Unmodified traditional MOs were used to block pcdh17 expression in the entire embryo from 1-4 cells stage, while a pcdh17 vivo-MO was used to block pcdh17 expression in the eye of older embryos (25-72 hpf).
Materials and methods
Morpholino design and microinjections. A translation blocking zebrafish pcdh17
MO (pcdh17atgMO: 5’-TGC ATC CCT TTC AGT GAG AGT GCC T-3’) which binds pcdh17 translation initiation region and inhibit the translation, and a pcdh17 splice- blocking MO (pcdh17sMO: 5’-ATA TAA GTT GTC GCT CCT ACC TGT A-3’) which bind exon1 and intron1 boundary (splice site) and lead to altered splicing of pcdh17 (e.g. inclusion of intron1, leading to premature stop of translation, see Fig. 7), were used to reduce pcdh17 function. For controls, a 5-base pair mismatch MO (5-misMO, 5’- ATA aAA cTT cTC cCT ACC TcT A-3’) was used for the splice-blocking MO, and uninjected embryos (from the same breeding parents) were used as blank control as well. All of the 18 three MOs were designed by and purchased from Gene Tools (Philomath, OR). MOs (one of the above) were injected (2-8 ng/embryo) into blastomere(s) or yolk immediately underneath the blastomere(s) of 1-4-cell stage embryos using a Narishige IM-300 microinjector (East Meadow, NY) under a stereo dissecting microscope. After microinjections, the embryos were placed in E3 medium and filtered fish tank water (1:1 volume), and allowed to develop at 28.5°C, to desired stages (e.g. 24, 36, 50 and 72 hpf, separately). The embryos or larvae were anesthetized and euthanized in 0.02% MS-222, and fixed in 4% paraformaldehyde. The embryos for WISH were processed as described above.
Reverse Transcriptase-Polymerase Chain Reaction. To demonstrate the efficacy of the pcdh17sMO which was designed to alter splicing of pcdh17, RT-PCR was used to amplify a pcdh17 fragment, using a pair of pcdh17 primers (P1 in Fig. 7, forward primer
5'-TGGTGCAAAGTGAAGTGGAG-3'; reverse primer 5'-
GGTGGAGGGTTACAAAGAAGC-3'), corresponding to nucleotides 2222 in zebrafish pcdh17 exon 1 to 45 nucleotides into the intron 1. For the same cDNAs from the control embryos and morphants, another pair of control primers bracketing part of zebrafish pcdh17 exon 1 (P2 in Fig. 7, nucleotides 1506-1958) were used. The latter amplified fragment was used as loading control on gel electrophoresis.
Whole mount immunocytochemistry (WICC). Fixed embryos were rehydrated in decreasing concentrations of methanol in PBST (PBS supplemented with 0.1% Tween
20). The embryos were treated with proteinase K (2 ug/ml in PBST), re-fixed in 4% paraformaldehyde, washed in PBST, and then PBST with supplements (100 mM PBS
19 with 1 mM DMSO, 0.5 mM 10% Tween 20). The embryos were incubated in blocking solution (3% normal horse serum in PBST with supplements) for 2 hours at room temperature, followed by incubation in a primary antibody solution (overnight on rotator at 4C) and then washed with PBST with supplements. Differentiating retinal cells or precursors could be labeled and distinguished by using different primary antibodies.
Primary antibodies used in this study included zn5 (Zebrafish Resource Center,
University of Oregon, stains axons and cell bodies of RGCs, 1:500 for fluorescent method and 1:1000 for peroxidase method), anti-HuC/HuD (Invitrogen, labels both
RGCs and amacrine cells, 1:1,100 and 1:2,000 for fluorescent and peroxidase methods, respectively), and zpr-1 (CHEMICON international, labels red-green double cone photoreceptors, 1:300 and 1:500 for fluorescent and peroxidase methods, respectively,
Endo et al., 1986; Macdonald and Wilson, 1997; Malicki, 1999; Malicki et al., 2003;
Masai et al., 2003; Liu et al, 2008). After washing the embryos with PBST, a secondary antibody was added to the embryos. For fluorescent WICC, the secondary antibody
(Jackson Laboratories, Bar Harbor, Maine) was labeled with Cy3 or FITC. After overnight incubation in dark on rotator at 4oC, the embryos were washed in PBST, kept in a VECTSHIELD mounting media (Vector Laboratories, Burlingame, CA) and observed via fluorescence microscopy. For peroxidase method, the secondary antibody was a biotylinated anti-mouse IgG (Vector Laboratories) and embryos were incubated in this secondary antibody solution overnight on a rotator at 4C. After washing in PBST, the embryos were placed in an avidin and biotin complex solution (Vector Laboratories) for
20 4-5 hours at room temperature or overnight at 4oC. Visualization of labeling was achieved using a DAB kit (Vector Laboratories).
Tissue sectioning. After being fixed in 4% paraformaldehyde, some embryos
(pcdh17 morphants, injected or uninjected control embryos) were washed in PBS, followed by incubation in 20% sucrose (dissolved in PBS) overnight at 4oC. 20% sucrose was replaced by new solution containing 50% O.C.T. and 50% of 20% sucrose. Six to eight embryos were transferred to an embedding mold made from household aluminum foil, which was filled with fresh O.C.T./20% sucrose (1:1) solution. Embryos were oriented with their heads pointing to the bottom of the mold. Rapidly frozen by a mixture of dry ice and 95% ethanol, the block was wrapped and kept in a dissecting bio-bag in -
20oC for short-term storage or -80oC for long-term storage. The tissue blocks were cut
12-15 m using a cryostat. Tissue sections were collected and mounted onto precoated glass slides (Fisher Scientific, Pittsburgh, PA) and dried at room temperature for about 1 hour. Slides were stored at -20oC till used for immunocytochemistry (ICC).
Immunocytochemistry on tissue sections (ICC). I performed ICC on tissue sections according to procedures described by Barthel and Raymond (1990). Briefly, tissues which were previously removed from freezer and dried at room temperature, were rehydrated in PBS, and incubated in a blocking solution for 40 minutes in a humid box to reduce non-specific staining. For 10 ml of PBS, I added 3 drops of normal goat serum
(ICC kit from Vector Laboratories), and 250 µl of 10% Triton X-100 (Sigma). The tissue sections were then incubated with a primary antibody dissolved in the blocking solution at 4°C overnight. Besides the antibodies mentioned in WICC, additional primary
21 antibodies were used for ICC on tissue sections: anti-beta-catenin (Sigma, St. Louis, MO; labels cell membranes and synaptic layers in embryonic zebrafish retina; 1:500; Liu et al.,
2008), and anti-Pax6 (Chemicon International, labels amacrine cells and precursors of horizontal cells, 1:500 for fluorescent method). After overnight incubation, unbound primary antibody was washed off in PBS at room temperature for 30 min. on a shaker.
Then tissues were incubated in a secondary antibody solution. For immunofluorescence, the tissue sections were treated with the secondary antibody conjugated to Cy3 or FITC
(either goat anti-mouse IgG or goat anti-rabbit IgG, Jackson Laboratories, 1: 100 in the blocking solution) for 2 hours at room temperature in dark, followed by washing in PBS
(30 min. at room temperature) and coverslipped in the VECTASHIELD mounting medium (Vector Laboratories). For the immunoperoxidase method, tissue sections were incubated with the biotinylated secondary antibody (ICC kit from Vector Laboratories,
1:250 in the blocking solution) at 4C overnight, followed by washing in PBS, and incubating in the avidin and biotin complex solution (ICC kit from Vector Laboratories) for two hours at room temperature. Visualization of this reaction was achieved using the
DAB kit (Vector Laboratories). After a satisfactory color reaction was achieved, PBS was applied to terminate the reaction. A few drops of glycerol were applied to each slide, and the slides were coverslipped, edges sealed with Permount (Sigma). The finished tissue slides were stored at -20oC till microscopy.
Analysis of retinal cell proliferation and cell death. To determine retinal cell proliferation, immunofluorescent ICC on retinal tissue sections was performed. Anti- phosphorylated histone H3 (PH3) antibody was used as the primary antibody (1:500,
22 Chemicon International) to label retinal cells in the M-late G2 phase of the cell cycle.
Numbers of proliferative cells from each embryo processed for the histone H3 immunostaining was obtained from three alternate sections from central retina (to avoid counting the same positive cells twice). Data collected were averaged to produce a value for that retina, using size and presence of the lens as reference points. Due to different eye sizes, numbers of histone H3-positive cells per unit eye area (100 μm2) were used for comparison. To determine apoptosis, terminal UTP nick-end labeling (TUNEL) was performed, according to the manufacturer’s instructions, on whole mount embryos using the Roche in situ cell death detection kit (Roche Molecular Biochemicals, Labat-Moleur, et al., 1998; Liu et al., 2008). Quantitative data of apoptotic cells were obtained from entire embryo eye of each whole mount embryo.
Microscopy and photography. Morphology of treated retina was observed as described before. To quantify eye sizes (surface area in square microns), 5 embryos from each stage stages (24, 50 and 72 hpf) were analyzed by SPOT Basic Software (SPOT imaging Solutions, Sterling Heights, MI). Unpaired Student’s t-test was used to determine statistical significance.
Vivo-pcdh17sMO application. Vivo-MOs are novel MOs covalently linked to a molecular scaffold that carries a guanidinium headgroup at each of its eight tips. This headgroup helps to remove the barrier of MOs’ entry into cells so that vivo-MOs could easily be applied to older embryos or adult organisms. A vivo-pcdh17sMO (sequence the same as the unmodified splice-blocking pcdh17MO) was injected into the right eyes of zebrafish embryos at 25-28 hpf, with the left eye as an internal control. Before the
23 injection, each embryo was anesthetized in tricaine methane sulfonate (MS222, Sigma, St.
Louis, USA), placed in a drop of 1% low-melting agarose gel on a glass slide. Another group of control embryos had their right eyes injected with a phenol red solution (20% in sterile water). After the injection, the embryos were freed from the agarose gel drops, and allowed to develop to specific stages (e.g. 50 and 72 hpf), and processed as described above. Then gross morphology, size, and differentiation of selective retinal cells of injected eye (right) were examined and internally compared to the control uninjected eyes
(left) using approaches described above. The embryos or larvae were euthanized in
MS222, fixed and processed for WICC using zn5 antibody. Some embryos were processed for cryosectioning and ICC on tissue sections using anti-HuC/HuD and zpr-1 antibodies.
Results
Injection of either the translation blocking pcdh17 MO (pcdh17atgMO) or splice- blocking pcdh17 MO (pcdh17sMO) into one- to four-cell stage zebrafish embryos
(pcdh17 morphants) resulted in embryos that had similar body shape and size as uninjected control embryos or embryos injected with the 5-mismatched pcdh17 MO, but with obviously reduced eye size (Figs. 5 and 6). The control eye from an uninjected embryo (Fig. 6A) was indistinguishable in appearance and size from the eye of an embryo injected with the 5-mis control MO (Fig 6B). Both were apparently larger than embryos injected with pcdh17sMO (Fig. 6C) or those injected with pcdh17atgMO (Fig.
6D), with the latter two similar in gross morphology and eye size (Fig. 6C and D).
24
Figure 5. Gross morphological defects in pcdh17 morphants were mainly confined to the eye. All images show lateral views of live embryos with anterior to the left and dorsal up. Eyes of embryos were indicated by asterisks. Scale bar = 200 μm for all images.
Figure 6. Higher magnification of eye regions of embryos in Fig. 3. All panels show lateral views (anterior to the left and dorsal up) of the head region from live embryos focusing on the eye. Scale bar = 100 μm for all images.
Efficacy of the pcdh17sMO was demonstrated by RT-PCR experiment showing that the morphants contained much higher levels of the abnormally processed mRNA
(inclusion of intron 1, Morcos, 2007) in the mRNA leading to a premature stop codon
123 nucleotides downstream of the exon 1, while only a faint band representing the
25 abnormally processed mRNA was observed in control embryos or adult brain tissue (Fig.
7). This is likely due to unspliced or aberrantly spliced transcripts in the control embryos
(Emond et al., 2009).
Figure 7. Diagnostic RT-PCR demonstrating efficacy of the splice-blocking pcdh17 MO. Lane 1 is 1 kb DNA ladder. Lane 2 (morphants cDNA) shows a PCR band of the correct size due to inclusion of intron 1 using the primer pair P1. Lane 3 (the same morphant cDNA as used in lane 2) is a PCR product using the primers pair P2. Lanes 4 (control embryo cDNA) and 5 (adult brain cDNA) show a faint band (asterisk) using the primer pair P1, likely due to unspliced or aberrantly spliced transcripts. Lanes 6 (control embryo cDNA) and 7 (adult brain cDNA) are loading controls using the primer pair P2.
26 Injection of different dosages of the pcdh17 MOs resulted in embryos with various degrees of defects. Slight to moderate defects (e.g. small eyes but no noticeable body defects) was detected in low-dose injected (1.5 ng/embryo) embryos, moderate to severe phenotypes (smaller eyes and heads, slightly smaller and/or curved bodies) were resulted from higher amount (3.0 ng/embryo) of injectionS (20.6% and 12.9% for pcdh17sMO and pcdh17atgMO, respectively, Table 1). The gross morphological defects were not obvious at 24 hpf, but became apparent at 50 hpf. To make analysis and interpretation of results more consistent, eye development and retinal cell differentiation were examined in moderately affected embryos injected with either pcdh17sMO or pcdh17atgMO.
Measurements of the eye size (area in square microns) of live embryos at 49 hpf and 72 hpf revealed that pcdh17 morphant eyes were significantly smaller (p<0.001) than uninjected control embryo eyes (Table 2). Moreover, embryos injected with the control 5- mismatched MO had similar eye size as the uninjected control embryos, while embryos injected with the pcdh17sMO had similar eye size as those injected with the pcdh17atgMO.
27
Table 1. Effects of pcdh17 MOs injections on zebrafish development
Number of Number of Number of Number of embryos with embryos with embryos with embryo
slight gross moderate gross severe gross examined at defects defects (%) defects (%) 48-50 hpf Uninjected 6* (2.0%) 0 2*(0.6%) 298 control pcdh17sMO 84 (41.0%) 89 (43.4%) 16 (7.8%) 205 (1.5 ng) pcdh17sMO 33 (11.5%) 187 (65.4%) 59 (20.6%) 286 (3.0 ng) pcdh17atgMO 39 (31.0%) 35 (27.8%) 7 (5.6%) 126 (1.5 ng) pcdh17atgMO 32 (14.8%) 124 (57.4%) 28 (12.9%) 216 (3.0 ng) 5-mis-sMO (3.0 5* (2.8%) 3* (1.7%) 3*(1.7%) 178 ng)
Gross morphological defects in the morphants are mainly seen in the eye (i.e. eye size). *The defects (e.g. smaller, curled, truncated bodies and/or much smaller or no heads and eyes) are different from those of morphants. All embryos are 48-50 hpf.
28 Table 2. Effects of pcdh17 MOs injections on eye size (region area in μm2)
Uninjected control pcdh17sMO pcdh17atgMO 5-mis-sMO 49 hpf 49,872 ± 3,825 33,096 ± 2,374* 33,500 ± 2,825* 49,740 ± 2,196 72 hpf 57,301 ± 2,469 43,875 ± 3,082* 43,591 ± 2,905* no data n=10 eyes for each group of embryos. Measurements were taken from the lateral side of the eye. *Using unpaired student t-test, the morphant eyes are significantly smaller (p<0.01) than either the uninjected control embryo eyes or embryos injected with the 5- bp mismatched MO. The eyes of 72 hpf morphants are significantly smaller (p<0.01) than those of 49 hpf control embryos.
Differentiation of retinal ganglion cells (RGCs) was examined at 49 hpf using zn5 antibody (labeling both RGC body and axons, Malicki et al., 2003; Masai et al.,
2003). A well-labeled RGC layer and optic nerve were seen in the retina of control embryos (Fig. 8A) or 5-mismatched MO injected embryos (Fig. 8B). In these embryos each optic nerve exited the retina, crossed the other optic nerve at the base of the diencephalon and projected toward their brain targets (Fig. 8A and B). Similar zn5 labeling pattern was observed in pcdh17 morphants (Fig. 8C and D; Table 3), except that the RGC layer was much reduced in size, and the optic nerves were much thinner compared to the control or 5-mismatched MO injected ones. Reduced RGC layer in pcdh17 morphants was confirmed using anti-HuC/HuD (labeling RGC body, Table 3,
Malicki et al., 2003; Masai et al., 2003). Differentiation of retinal cells in 72 hpf pcdh17 morphants was examined by anti-HuC/HuD (labeling soma of both RGCs and amacrine cells at this stage) and zpr-1 (labeling double cones) immunostaining (Fig. 8E-H, Table 3,
Malicki et al., 2003; Masai et al., 2003). In control embryos (Fig. 8E) or embryos 29 injected with the 5-mismatched MO (Fig. 8F), there were well-laminated RGC layer, amacrine cell layer and photoreceptor layer. Moreover, in these embryos there was a distinct unlabeled gap between the RGC layer and the amacrine cell layer, where RGCs, amacrine cells and bipolar cells synapse (ipl). In pcdh17 morphant retina (Fig. 8G and H,
Table 3), regions expressing those markers, especially zpr-1, were much reduced, and ipl was barely detectable. To further examine pcdh17 function on retinal lamination, I used
β-catenin immunostaining (labeling cell membrane of retinal cells and retinal synaptic layers) and showed that retinal layer organization was not much affected in most 72 hpf pcdh17 morphants, but ipl was much reduced compared to that in control embryos (Fig.
9, Table 3).
Figure 8. Immunostaining analysis of retinal cell differentiation in control animals (panels A and E), animals injected with the 5-miss MO (panels B and F), and pcdh17
30 morphants (panels C, D, G and H). Panels in the top row showed ventral views (anterior up) of the head region of 49 hpf whole mount embryos processed for zn5 peroxidase immunostaining, while panels in the bottom row showed cross retinal sections (dorsal to the left, from central retina) from 72 hpf embryos labeled with anti-HuC/D and zpr-1 antibodies (immunofluorescent methods). Arrows in panels F and G indicated location of the optic nerve. Abbreviations: ipl, inner plexiform layer; on, optic nerve; te, telencephalon. Other abbreviations are the same as above. Scale bars = 100 and 50 µm for the top and bottom panels, respectively.
Figure 9. Retinal organization of a control embryo (panel A) and a pcdh17 morphant (panel B) revealed by beta-catenin immunostaining. The images are from retinal cross sections (dorsal to the left) labeled with a beta-catenin antibody. Abbreviations: opl, outer plexiform layer. Other abbreviations are the same as above. Scale bar = 50 µm.
31 Table 3. Effects of pcdh17 knockdown on zebrafish retinal development revealed by immunostaining
Hu Hu&zpr-1 zn5 beta-catenin 48-50 hpf Uninjected 0% (n=20) ND 12.5% (n=40) ND control 100% pcdh17sMO ND 100% (n=30) ND (n=20) pcdh17atgMO ND ND 100% (n=24) ND 5-mis .3% (n=24) ND 13.3% (n=30) ND pcdh17sMO
72 hpf Uninjected ND 8.0% (n=25) ND 0% (n=6) control pcdh17sMO ND 85% (n=20) ND 16.7% (n=6) pcdh17atgMO ND 100% (n=12) ND ND 5-mis ND 0% (n=8) ND ND pcdh17sMO n, number eyes examined; %, percentages of greatly reduced or altered staining staining (staining area and/or intensity) compared to the majority of control embryos. Abbreviations: Hu, anti-HuC/HuD immunostaining; Hu & zpr-1, anti-HuC/HuD and zpr-1 double immunostaining; ND, analysis not done; zn5, zn5 immunostaining. (line space reduced)
The small eyes in pcdh17 morphants could be due either to increased cell death as in cad4 morphants retinae (Babb et al., 2005) or reduced cell proliferation as in cad6 morphants retinae (Liu et al., 2008). TUNEL labeling was performed to examine 32 apoptosis in whole mount embryos (Fig. 10), while histone H3 immunostaining was used to determine retinal cell proliferation (Fig. 11, Adams et al., 2001). There were very few apoptotic cells in the retina of both young (25 hpf, one TUNEL-positive cell in twelve eyes) and older (50 hpf, three TUNEL-positive cells in twenty-two eyes) pcdh17 morphants (Fig. 10D and E), which was similar to uninjected control embryos (Fig. 10A and B, one TUNEL-positive cell in twelve eyes at 25 hpf, four TUNEL- positive eyes in twenty-six eyes at 50 hpf). These low numbers of TUNEL-positive cells in the eye were unlikely due to experimental errors, because numerous labeled cells were detected in the trunk and tail regions of these embryos (Fig. 10C and F).
Figure 10. Apoptosis analysis using TUNEL staining. All panels are lateral views of whole mount embryos with anterior to the left and dorsal up. Panels A, B, D and E are lateral views of the head region focusing on the eye, while panels C and F are from posterior trunk and tail regions of the same embryos shown in panels B and E, respectively. Abbreviations are the same as above. Scale bars = 100 μm for all panels.
33 Mitotic nuclei in the control and pcdh17 morphant retinae were revealed using the histone H3 immunostaining (Fig. 11). As in the retina of control embryos (Fig. 11A), most labeled cells in morphant retina (Fig. 11B) were detected in the peripheral region at
50 hpf. But there were significantly more histone H3 positive cells in the control retina than pcdh17 morphant retina (Fig. 11C).
Figure 11. Histone-H3 immunostaining in control retina (panel A) and pcdh17sMO morphant retina (panel B). The images are cross sections (10 μm) from central retina (dorsal to the right). Retinal areas are outlined by the dashed line. Arrows indicate a few labeled nuclei in focus. The data in panel C is from 20 eyes (10 for each group). Asterisks indicate highly significant differences (p<0.0001) between the control and morpholino treatment. Abbreviations are the same as above. Scale bar = 50 µm for panels A and B.
34 To assess a more direct pcdh17 function in the zebrafish eye development, I used vivo MO method, a newly developed MO technology described by Gene Tools
(pubs.gene-tools.com). Embryos injected with the vivo-MO had similar gross morphology as uninjected control embryos (Fig. 12A and B), but had smaller eyes compared to uninjected control embryos or embryos injected with phenol red (Fig. 12C-
E, Table 4). At 49 and 72 hpf, both the right eye (injected) and the left eye (uninjected) were significantly smaller than those of the control embryos, but the vivo-morphant right eye was significantly smaller than the left eye (Fig. 12 and Table 4).
Figure 12. Gross body (panels A and B) and eye (panels C-E) morphology in live control and vivo-morphant embryos. Panels A and B show lateral views (anterior to the right and dorsal up) of the right side of a control embryo (panel A) and vivo-morphant (panel B). Panels C-E show ventral views (anterior to the top) of the head region of a phenol red injected embryo, and two vivo-pcdh17sMO injected embryos. Injections were applied to the right eye of the embryos. Defects in panel E was more severe than in panel D. Scale bars = 250 µm for panels A and B, and 50 µm for panels C-E.
35 Table 4. Effects of vivo-pcdh17sMO injection on eye size
Uninjected control injected control vivo-pcdh17sMO 49 hpf Left eye 49,858 ± 961 (n=6) ND 45,107 ± 2,433 (n=12)* Right eye 50,358 ± 1,534 (n=6) ND 40,001 ± 2,561 (n=12)*
72 hpf 57,924 ± 2,420 Left eye ND 53,390 ± 3,555 (n=12)* (n=12) 58,763 ± 1,701 Right eye ND 46,098 ± 2,838 (n=12)* (n=12)
Region area in μm2. n, number of eyes examined. Measurements were taken from the lateral side of the eye. *The vivo-morphant eyes are significantly smaller (p<0.01) than either the uninjected control embryo eyes or phenol red injected (only to the right eye) embryo eyes.
Zn5 immunostaining revealed that labeling of the RGC layer and/or the optic nerve was obviously changed (i.e. smaller zn5-positive RGC layer and/or thinner optic nerve) in less than half (43.4%) of the left eye (uninjected), but in the vast majority
(87.5%) of the right eye (injected) of the vivo-morphant (Fig. 13A-C, Table 5) at 49 hpf.
At 72 hpf, similar percentages (41.6% and 91.7% for the left and right eye, respectively) of vivo-morphant eyes had altered anti-HuC/HuD and zpr-1 staining, with reduced immunoreactive regions in the RGC layer, amacrine cell layer, photoreceptor layer, and narrower ipl (Fig. 13D-I). As in the eye size and zn5 staining, the retinal defects in 72 hpf
36 vivo-morphants were more severe in the injected right eye (Fig. 13F and I) than the uninjected left eye (Fig. 13E and I).
Figure 13. Immunostaining analysis of retinal cell differentiation in vivo-morphant retina. Panels A-C are ventral views (dorsal up) of the head region of 49 hpf whole mount embryos processed for zn5 immunoperoxidase staining. Panels D-I show cross retinal sections (dorsal to the left, from central retina) of 72 hpf embryos processed for anti- HuC/D and zpr-1 immunofluorescent labeling. Arrows in panels D, E, G and I point to location of the optic nerve. Abbreviation: inj-control, phenol red injection control. Other abbreviations are the same as above. Scale bars = 100 µm for panels A-C, 50 µm for the remaining panels.
37 Table 5. Effects of vivo-pcdh17sMO injection on zebrafish retinal development revealed by immunostaining
Zn5 Hu&zpr-1 49 hpf Uninjected control 9.4% (n=32) ND Morphant left eye 43.4% (n=16) ND Morphant right eye 87.5% (n=16) ND Phenol red injected control 12.5% (n=16) ND
72 hpf Uninjected control ND 0% (n=10) Morphant left eye ND 41.6% (n=12) Morphant right eye ND 91.7% (n=12) Phenol red injected control ND 10.0% (n=10) n, number eyes examined; %, percentages of greatly reduced or altered staining (staining area and/or intensity) compared to the majority of control embryos, or reduced inner plexiform layer. Abbreviations: Hu & zpr-1, anti-HuC/HuD and zpr-1 double immunostaining; ND, analysis not done; zn5, zn5 immunostaining.
38 CHAPTER V
MOLECULAR MECHANISM UNDERLYING pcdh17 FUNCTION
Retinal regulatory molecules (crx, otx5, neuroD, and rx1) are a group of transcription factors that are considered essential for the vertebrate eye development.
Expression of most of these molecules was reduced in zebrafish retina with reduced classical cadherins function (Liu et al., 2007; 2008). I would like to know if their expression was also changed in the retina of pcdh17 morphants.
Proteins are functional elements of dynamic organisms (Honore et al., 2004).
Proteomics was used to reveal protein expression patterns under different developmental conditions: control and MO-injected. In this experiment, protein expression of the entire proteome between these two groups was compared. Analysis of the expression patterns would provid me with insights into protein signaling and related biology pathways.
Materials and methods
WISH of transcription factors. cDNAs used to generate the cRNA probes were generously provided by Pamela Raymond at the University of Michigan (for crx, otx5, and rx1 genes), and Deborah Stenkamp at the University of Idaho (for neuroD). The dig- labeled cRNA probes were used for assessing differentiation of pcdh17 morphants retinae.
The WISH was performed as described above.
39 Protein extraction and two-dimensional polyacrylamide-gel electrophoresis (2D
PAGE). Two-dimensional gel electrophoresis was used as the central tool for assaying the proteome. Embryos (30-60 per group) were collected at desired stage (50 and 72 hpf, separately), and ultrasonicated in 500 μl of 50 mM HEPES buffer (pH 7.8) on ice using three 3-s bursts with a 5-s rest between bursts (Sonic Dismembranator, Fisher Scientific,
PA). A Bradford protein assay was performed to measure protein concentration (Bradford,
1976). A Perfect-FOCUS kit (GenoTech, St. Louis, MO) was used to remove nucleic acids, salts, lipid and other non-protein substances following the manufacturer’s instructions. First dimension electrophoresis with immobilized pH gradient-isoelectric focusing (IPG-IEF) was performed following the standard protocol of Toka and Kimura
(1997). Briefly, IPG-IEF strips, (pH 3-10, Bio-Rad, Hercules, CA) were rehydrated overnight in IEF loading buffer (7M Urea, 2M Thiourea, 1% DTT, 1% CHAPS (3-([3- cholamidopropyl]-dimethylammonio-2-hydroxy-1-propanesulfonate), 1% Pharmalyte, 1%
Triton and trace bromophenol blue) (Active 50V, 20C). For silver nitrate stained gels,
100 µg total protein was loaded per strip (1 mg for Coomassie blue staining). The next day, strips were focused continuously at maximum current 50 mA until Volt Hours reached ~28000 VH. Strips were subsequently equilibrated for 10 min. in 6 M urea, 1%
SDS, 375 mM Tris–HCL (pH 8.8), 30% glycerol and 100 mM DTT, followed by another
10 min. in the same buffer with 450 mM iodoacetamide substituted for DTT and a trace bromophenol blue. After the second equilibration, strips were immediately loaded on the second dimension. Second dimension separation was undertaken on vertical 4-15% precast polyacrylamide gels (Bio-Rad, CA). Each equilibrated strip was placed in contact with the top of a gel. PINK plus prestained protein ladder (Insight Genomics, Falls
40 Church, VA) was used for molecular weight estimation. The gel was applied with 200 V and run at room temperature until the blue dye reached the gel bottom. The gel was subsequently fixed overnight (50% Ethanol, 10% Acetic Acid), and washed in distilled water three times, 10 min. each with agitation. Then the gels were processed to staining steps (Silver nitrate staining and Coomassie Blue staining).
Silver nitrate staining (sensitivity 1–10 ng). Each washed gel was incubated in 10% glutaraldehyde for 20 min., and washed again using distilled water for 2 hour, changing every 10 min. to reduce the background. Then, the gel was washed in 5 μg/ml DTT solution for 30 min. with agitation. Without rinsing, the gel was agitated in 0.1% silver nitrate solution for 30 min. after the DTT incubation. The gel was developed in a developing solution (500 ml developing solution: 0.9 ml of 10% formaldehyde, 15 g of sodium carbonate) until desired silver staining was achieved. A stop solution (5 ml of 2.3
M citric acid per 100 ml developing solution) was added to finalize the staining step.
Coomassie blue staining (sensitivity 100 ng). After finishing the fixation, the gel was washed and incubated in Coomassie blue G-250 stain Code solution (GelCode Blue,
Pierce, Rockford, IL) until desired staining was achieved. Uncombined blue dye was washed by distilled water.
Gel acquisition and analysis. Stained gels were scanned with a transmittance scanner (Epson 2450, 3.3 Dynamic Range) and analyzed by Melanie software (version
6.0, Genebio, Geneva, Switzerland). A reference gel was chosen to which all other gels were compared by matching two significant land markers as well as manual editing and verification of the matched spots. Inter-group gel matching was performed, and relative pixel volume (Vol%, (spot area*density)/total pixel density) was estimated. Differentially
41 expressed proteins between groups were defined in following ways: mean Vol% ≥ two- fold difference and/or spots presence in one group but absent in the other.
Protein Identification and Verification. After differentially expressed protein spots were chosen, they were excised from the gel and processed for Mass spectrometry (MS).
In-gel digestion was manually performed following a protocol detailed in Goel et al.,
2005. Selected spots were cut with a razor blade from the Coomassie blue stained gel.
Each spot was diced into 1 mm3-cubes and placed in a siliconized tube which pre-rinsed with ethanol (95% USP grade). Excised samples were washed and destained in 175 μl of wash reagent (50% ethanol with 5% acetic acid in distilled water) for 1 hour at RT twice.
Dehydration of samples was followed by adding 175 μl of acetonitrile (HPLC grade,
Burdick and Jackson, Morristown, NJ) for 5 min. and sample pieces were rehydrated by adding 175 μl of bicarb solution (100 mM of bicarb solution: 0.2 g ammonium bicarbonate in 20 ml distilled water) for 5 min. Dehydration by using acetonitrile was performed for another 5 min. and then the samples were dried in SpeedVac for 3 min. The samples were digested in 10 μl of 10 ng/μl sequencing grade modified trypsin (Promega,
Madison, WI) in 2000 μl of 50 mM bicarb solution overnight. The digested samples was then incubated in 30 μl of extraction reagent (50% of acetonitrile; 5% formic acid, ACS reagent grade, Fisher, Pittsburgh, PA) for 10 min. the following day. Gel pieces were removed and extracted solution was left. Another 30 μl of extraction reagent were added into each original tube. After mixing by vortex, volume of solution was reduced to less than 10 μl (totally dried is OK) in SpeedVac. Then samples were sent to Cleveland Clinic
Mass Spectrometry Laboratory to be analyzed. MS analysis of samples was performed by the Cleveland Clinic Mass Spectrometry Laboratory
42 (http://www.lerner.ccf.org/services/mass_spec/). Briefly, the digested samples were resuspended in 1% acetic acid and 10 μl of the extract were injected into the capillary high-performance liquid chromatography (HPLC) column (operator-packed 9 cm x 75
μm id Phenomenex Jupiter C18 reversed-phase). Peptides eluted from the column at a flow rate of 0.25 μl/min. by an acetonitrile/0.1% formic acid gradient and simultaneously introduced into a Finnigan LTQ linear ion trap mass spectrometer system (ThermoFisher,
San Jose, CA, USA) which had a 2.5 kV microelectrospray ion source. Peptide molecular weights were determined, and detailed amino acid sequence was further determined by collisionally induced dissociation (CID) spectra. Peptide peaks were submitted to Mascot
(www.matrixscience.com, Perkins et al., 1999) to obtain initial protein identification.
Parameters were set as described in Jury et al., 2008. The great majority (83%) of identifications were made by more than three peptides covering more than 10% of total amino acid sequence. After gaining protein identifications, proteins were searched in
Gene Ontology databases (GO, http://www.gengeontology.org/).
ICC confirmation. To confirm the results from MS, immunocytochemistry was performed as described in Chapter IV. Müller cell antibody zrf-1 (1:5,000 for fluorescent staining, Zebrafish International Resource Center, Eugene, OR) was used to confirm the down regulation of glial fibrillary acidic protein (GFAP) in morphants, as predicted from proteomics study.
Results
To determine whether pcdh17 regulates retinal differentiation through affecting expression of retinal regulatory molecules, four transcription factors known markers for
43 vertebrate retinal differentiation (Table 6) were examined in control and pcdh17 morphant retinae: rx1 (Chuang et al., 1999; Chuang and Raymond, 2001), NeuroD
(Morrow et al., 1999; Inoue et al., 2002; Yan et al., 2005), crx (Furukawa et al., 1997;
Blackshaw et al., 2001; Liu et al., 2001; Shen and Raymond, 2004) and otx5 (Gamse et al., 2002). Blocking pcdh17 function appeared to have little effect on early development of the eye as rx1 expression was similar between the control and morphant retinae (Fig.
14A and E, Table 6). neuroD, crx and otx5 were strongly expressed throughout the control retina at 49 hpf (Fig. 14B-D), but their expression was greatly reduced in pcdh17 morphant retina (Fig. 14F-H; Table 6). crx expression was especially diminished, almost absent in the posterior half of the morphant retina (Fig. 14G). In contrast, crx and otx5 expression in the morphant pineal gland was similar to control embryos (inserts in Fig.
14C, D, G and H).
44 Table 6. Effects of pcdh17 knockdown on expression of transcription factors and in the retina
rx1 neuroD Crx otx5 24 hpf Uninjected 0% ND ND ND control (n=24) 0% pcdh17sMO ND ND ND (n=16) 0% pcdh17atgMO ND ND ND (n=18)
48-50 hpf Uninjected ND 0% (n=16) 0% (n=14) 0% (n=20) control pcdh17sMO ND 100% (n=12) ND 100% (n=12) pcdh17atgMO ND 100% (n=12) 100% (n=16) 100% (n=14) 5- ND 0% (n=10) ND 0% (n=10) mispcdh17sMO n, number of eyes examined for each probe; %, percentages of greatly reduced or altered staining (staining area and/or staining intensity), compared to the majority of control embryos. Abbreviation: ND, analysis not done.
45
Figure 14. Expression of transcription factors in the control and pcdh17 morphant retinae. Embryos were processed for whole mount in situ hybridization. Panels A and E are in- face views (dorsal up) of embryo heads. The remaining panels are lateral views (anterior to the left and dorsal up) of eyes and/or heads. Arrows in the insert in panels C, D, G and H point to the pineal gland. Abbreviations are the same as above. Panels A and E are of the same magnification, while the remaining panels have the same magnification as panel B. Insert in panels C, D, G, H are of the same magnifications. Scale bars = 100 µm.
In both 50 hpf gels and 72 hpf gels, there were more expressed proteins (total number on individual spots) in the control group than the morphant group (Fig. 15). In addition, there were more spots on 50 hpf gels than 72 hpf gels for both the control (Fig.
15 A and C) and morphant groups (Fig. 15 B and D). However, a higher proportion of identified proteins were yolk proteins (i.e. isoforms of vitellogenin in 50 hpf gels).
Therefore, I chose 72 hpf embryos with smaller yolk size and higher percentage of non- vitellogenic proteins for further analysis.
46 Figure 15. Blue stained gels of total proteins of developing zebrafish. Upper panels were 50 hpf control (A) and morphant embryos (B). Panel C and D showed protein expression of 72 hpf control and morphant embryos, respectively. For each panel, left to right: pI 3- 10; top to bottom: MW 120-10 kDa.
Coomassie Blue stained gels of 72 hpf embryos are shown in Figure 16. The sum total of quantifiable spots was higher in the control group than the morphant group
(516±15 vs. 333±11, p<0.05). From my selection of significantly differently stained spots, three of them (Table 7, first part, SB13-26-23, SB13-26-24, SB13-26-25) were only shown in the control group; four of them (Table 7, second part, SB13-26-26, SB13-
26-27, SB13-35-3, SB13-35-4) were expressed at a lower relative values in the control group; five of them (Table 7, third part,SB13-26-28, SB13-35-5, SB13-35-6, SB13-35-8,
SB13-35-10) had decrease relative values in the morphant group.
47 Figure 16. Coomassie blue staining 2-DE electrophoresis map of developing zebrafish (72 hpf control group) total protein. All labeled spots were characterized by MS. Ladder on the top is pH value (3 to 10); ladder on the left is ladder of Molecular Weight (MW, kD).
48 Table 7. Identification parameters for zebrafish MS analyzed protein bands (showed in Fig. 16)
Sampl M AVG AVG Rati Protein name NCBI IN pI % e # W Con. MO o 4 Muscle creatine kinase a 18858427 43 6.3 5 15778718 2 SB Muscle creatine kinase b 43 6.3 1 7 13262 1.36 0 NA Phosphoglycerate kinase 1 3 47087077 45 6.5 1 7 1 Beta-actin-like protein 2 50344802 42 5.2 0 15778718 4 SB Muscle creatine kinase b 43 6.3 1 8 13262 1.04 0 N/A 1 4 Muscle creatine kinase a 18858427 43 6.3 9 SB Actin, alpha cardiac 4 13262 18858249 42 5.2 0.88 0 N/A muscle 1 4 5 Ubiquitin-conjugating 1 SB 41054401 17 5.7 enzyme E2Nb 8 13262 0.18 0.54 0.33 Ubiquitin-conjugating 1 6 47087307 17 5.7 enzyme E2Na 8 Histone H2A type 1-A- 29260936 10. SB 15 11 like isoform 1 8 4 13262 0.16 0.45 0.36 Tubulin-specific 2 7 41054980 13 5.4 chaperone A 8 SB 60 kDa heat shock 31044489 2 61 5.6 0.04 0.09 0.44 13353 protein, mitochondrial 1
49 SB 78 kDa glucose- 2 47085775 72 5.0 0.06 0.14 0.43 13354 regulated protein 8 40S ribosomal protein 6 SB 41152459 15 6.8 S12 1 13262 0.34 0.12 2.83 5 8 Histone H4-like 68362808 11 11.4 0 SB prohibitin 41152028 29 5.3 3 0.27 0.02 13.5 13355 29262730 4 Profilin-1 isoform 1 15 6.6 SB 5 0 0.15 0.03 5 13356 3 Histone H4-like 68362808 11 11.4 8 4 Keratin 12 51010971 50 5.3 0 Glial fibrillary acidic 3 SB 66393075 51 5.3 protein 0 0.16 0.03 5.33 13358 Keratin 4 18858947 54 5.4 9 Protein disulfide- 1 41282163 48 5.2 isomerase A6 7 Triosephosphate 3 SB 47271422 27 6.9 isomerase B 4 13351 0.23 0.06 2.83 Hypothetical protein 2 0 62955557 27 7.0 LOC550490 0
Sample #: Spot number in Fig. 14.
NCBI IN: NCBI database index number.
MW: Calculated molecular mass, in kDa. pI: Isoelectric point.
%: Percentage of the complete primary sequence covered by matching peptides.
50 AVG Con.: Mean relative pixel volume for control groups.
AVG MO: Mean relative pixel volume for morphant groups.
Ratio: The ration of mean relative pixel volume of control groups and morphant groups. Only more than 2 or less than 0.5 folds were selected to do MS analysis.
51 GO terms were searched to determine which cellular function and biological process responded to all differentially expressed proteins. Some of differentially expressed proteins in Table 7 were uncharacterized in Zebrafish database: Beta-actin-like protein 2, Histone H2A type 1-A-like isoform 1, 78 kDa glucose-regulated protein,
Histone H4-like, Keratin 12, hypothetical protein LOC550490. Others were annotated and details were listed in Table 8, 9 and 10.
Proteins with high quality annotations included structural proteins (actin), heat shock proteins (60-kDa HSP), molecular chaperone proteins (Tubulin-specific chaperone
A), cell cycle proteins (prohibitin), and enzymes (creatine kinase, skeletal muscle). The decrease in the muscle-specific creatine kinase indicates less energy production and lower metabolism in the morphant group. Increased tubulin-specific chaperone A in the morphants suggests that correctly folding process of functional tubulin was inhibited since this cofactor controls tubulin capture and stabilization during tubulin folding (Tian et al., 1996). Heat shock protein 60, which is involved in cell cycle control, was more abundant in the control embryos, suggesting higher nuclear β-catenin transcriptional activity in the morphant (Tsai et al., 2009). Nuclear histone protein (H4 like), with decreased expression in the morphant, is implicated in the Notch signal pathway.
Increased expression of ubiquitin-conjugating enzymes in the morphant group suggests higher protein turnover. Glial fibrillary acidic protein (GFAP), a marker for developing of
Müller glia, was down-regulated in the morphants, suggesting its underlying Notch-Delta signaling pathway was negatively impacted by MO injection (Bernardos et al., 2004).
52 Table 8. Gene ontology Biological Process Terms of the MS identified proteins
GO:0008152 metabolic process GO:0044267 cellular protein metabolic process GO:0016310 phosphorylation GO:0006412 translation GO:0006950 response to stress GO:0006457 protein folding GO:0006096 glycolysis GO:0006098 pentose-phosphate shunt GO:0006094 gluconeogenesis GO:0007010 cytoskeleton organization GO:0006334 nucleosome assembly GO:0042026 protein refolding GO:0030036 actin cytoskeleton organization GO:0000724 double-strand break repair via homologous recombination GO:0001666 response to hypoxia GO:0000209 protein polyubiquitination GO:0043123 positive regulation of I-kappaB kinase/NF-kappaB cascade GO:0051092 positive regulation of NF-kappaB transcription factor activity GO:0042246 tissue regeneration GO:0031058 positive regulation of histone modification GO:0050852 T cell receptor signaling pathway GO:0016574 histone ubiquitination GO:0006301 postreplication repair GO:0007021 tubulin complex assembly GO:0045739 positive regulation of DNA repair GO:0051443 positive regulation of ubiquitin-protein ligase activity GO:0060027 convergent extension involved in gastrulation GO:0000729 DNA double-strand break processing GO:0031101 fin regeneration GO:0055113 epiboly involved in gastrulation with mouth forming second GO:0042769 DNA damage response, detection of DNA damage GO:0033182 regulation of histone ubiquitination
53 Table 9. Gene ontology Molecular Function Terms of the MS identified proteins GO:0003824 catalytic activity GO:0016740 transferase activity GO:0000166 nucleotide binding GO:0016772 transferase activity, transferring phosphorus-containing groups GO:0005524 ATP binding GO:0003677 DNA binding GO:0016301 kinase activity GO:0016874 ligase activity GO:0016853 isomerase activity GO:0005198 structural molecule activity GO:0003735 structural constituent of ribosome GO:0016881 acid-amino acid ligase activity GO:0051082 unfolded protein binding GO:0003779 actin binding GO:0004618 phosphoglycerate kinase activity GO:0004807 triose-phosphate isomerase activity GO:0043130 ubiquitin binding GO:0003756 protein disulfide isomerase activity GO:0017072 tubulin-specific chaperone activity
54 Table 10. Gene ontology Cellular Component Terms of the MS identified proteins GO:0005575 cellular_component GO:0016020 membrane GO:0005622 intracellular GO:0005737 cytoplasm GO:0030529 ribonucleoprotein complex GO:0005840 ribosome GO:0005634 nucleus GO:0005694 chromosome GO:0005856 cytoskeleton GO:0000786 nucleosome GO:0015629 actin cytoskeleton GO:0005882 intermediate filament GO:0005759 mitochondrial matrix GO:0045095 keratin filament GO:0031372 UBC13-MMS2 complex GO:0045098 type III intermediate filament
Immunocytochemistry on cryosectioned tissues was performed to independently confirm results from the proteomics study. Zrf-1 was used as the primary antibody
(Zebrafish International Resource Center, Eugene, OR). This monoclonal antibody labels
Müller radial glial cells in embryonic zebrafish (Trevarrow et al., 1990). In the retina of control embryos (Fig. 17A), well-labeled Müller glia cells were seen spanning almost the entire thickness of the retina, while the staining was greatly reduced in the morphant retina (Fig. 17C), and the labeling was confined mainly to 1/3 the thickness of the retina
(i.e. the retinal ganglion cell layer). In contrast, GFAP labeling was similar in the mid/hind brain region between the control (Fig. 17B) and morphant embryos (Fig. 17D).
55
Figure 17. Immunohistochemstry confirmation of GFAP expression in the control (upper panels) and morphant tissues (lower panels). All panels show cross sections. Panels A and C have dorsal to the left, while panels B and D have dorsal up. GFAP labeling is indicated by the arrow in panels A and C. Abbreviations: hy, hypothalamus; ot, optic tectum; teg, tegmentum. Other abbreviations are the same as Fig. 6. Panel A and C are of the same magnification, while panels B and D are of the same magnification. Scale bar = 50 µm.
56 CHAPTER VI
DISCUSSION AND CONCLUSIONS
Pcdh17 is essential in zebrafish eye development
pcdh17 expression in the zebrafish retina begins about 26 hpf in the anteroventral region (see Chapter III) where retinal cell differentiation is initiated (Nawrocki, 1985; Hu and Easter, 1999). Retinal ganglion cells (RGCs) are the first retinal cells to differentiate in this region at about 28 hpf (Nawrocki, 1985). This temporal and spatial correlation between the RGCs differentiation and pcdh17 expression suggests that pcdh17 plays a role in the initial differentiation of the retinal ganglion cells. pcdh17 is expressed in the entire retina at 34-50 hpf (see Chapter III) when retinal ganglion cells continue to differentiate as their earliest axons grow out of the retina, pass the optic chiasm (32-34 hpf), and project toward their brain targets, while the second type of retinal cells, the amacrine cells, starts to differentiate by 40 hpf (see Literature reviews). Pcdh17 therefore likely continues to participate in the RGCs differentiation (e.g. by promoting retinal axons outgrowth and elongation), and also is likely involved in the initial and continued a differentiation of the amacrine cells. The embryonic retina continues to express pcdh17 at 50-72 hpf, with decreased expression in the inner half of the retina where retinal ganglion cells and amacrine cells reside, while increased expression in the outer half of
57 the inner nuclear layer where the horizontal cells are located (see Chapter III and
Literature reviews). This expression pattern suggests that pcdh17 is less involved in the late differentiation of the RGCs and amacrine cells, but plays a role in the differentiation of the horizontal cells (begins around 57 hpf).
There are numerous studies on cadherins expression in the vertebrate retina (see
Literature reviews). Several cadherins, such as cadherin-2 and pcdh19, are also expressed in the entire zebrafish retina, but their expression start earlier (before 24 hpf) and more or less remain unchanged during the first three days of development (see
Literature reviews). My results provide further supporting evidence that each cadherin has a unique spatial and/or temporal expression pattern.
Results from my subsequent functional study confirm the idea that pcdh17 plays an important role in zebrafish retinal cells differentiation. Disrupted development of
RGCs, amacrine cells and photoreceptors in pcdh17 morphants suggests that pcdh17 participates in differentiation of these cells. Moreover, pcdh17 may be more directly involved in RGCs and amacrine cells differentiation than the photoreceptor development, because pcdh17 is expressed throughout the retina of 34-72 hpf, except in the outer nuclear layer (photoreceptors reside) where its expression is weaker at 50 hpf (Fig. 4C) when this layer is becoming distinguishable, and its expression is more reduced at 72 hpf
(Fig. 4D). Reduced photoreceptors differentiation may, at least partially, due to pcdh17 effect on differentiation of retinal cells located in the inner nuclear layer such as the horizontal cells (located in the outer portion of the inner nuclear layer where pcdh17 expression remains strong at 50-72 hpf), which then affects photoreceptor development.
Horizontal cells differentiation was not examined in pcdh17 morphants mainly due to
58 lack of reliable horizontal cell markers. It is also possible that pcdh17 is directly involved in the development of the earliest formed photoreceptors (e.g. blue cones and double cones) that begin to express photoreceptor specific markers as early as 40-44 hpf, but is less involved in differentiation of later formed photoreceptors (e.g. red cones and rods that start to form between 48 and 60 hpf, Larison and Bremiller, 1990; Robinson et al.,
1995; Schmitt and Dowling, 1996; Raymond et al., 1996). The smaller eye phenotype in pcdh17 morphants is likely due mainly to reduced retinal cell proliferation, which is similar to Xenopus with reduced cdh6 function (Ruan et al., 2006), and zebrafish cad6 morphants (Liu et al., 2008).
Compared to zebrafish embryos with disrupted classical cadherins expression
(e.g. cdh2, cdh4 and cdh6, Lele et al., 2002; Malicki et al., 2003; Masai et al., 2003;
Babb et al., 2005; Liu et al., 2008), pcdh17 morphants show defects that are mainly confined to the eye, and less severe retinal defects. Despite similar cdh2 and pcdh17 expression (throughout the retina) in the retina of 34-50 hpf embryos, cdh2 morphants and mutants showed disrupted retinal lamination (Malicki et al., 2003; Masai et al.,
2003), while pcdh17 morphants have normal appearing retinal lamination. It is possible that cdh2 compensates for pcdh17 function in retinal lamina formation, or later expressed molecules (e.g. pcdh17 see above) have less effect on the formation of the retinal lamination than earlier expressed molecules (e.g. cdh2 throughout the retina at 24 hpf,
Liu et al., 2001). Although cdh4 and cdh6 are expressed later (after 30 hpf) and have more restricted expression (confined mainly to the inner portion of inl, Liu et al., 1999;
2001), differentiation of RGCs, amacrine cells and photoreceptors is much more severely affected in these morphants (Babb et al., 2005; Liu et al., 2008) than in pcdh17
59 morphants. This may partially due to wider expression of these classical cadherins in other parts of the CNS and/or other tissues. For example, cdh6 is expressed in both the
CNS and selective mesodermal structures (e.g. cartilages, kidney). Embryos with disrupted cdh6 function also show apparent head and body defects (e.g. enlarged pericardial cavity and abnormally developed kidney, Kubota et al., 2007; Liu et al.,
2008). Again, the less severe retinal defects observed in the pcdh17 morphants may due to compensatory function of other cadherins (e.g. cadherin-2, -4 and -6) expressed in the developing retina.
Loss-of-function techniques
Conventional MOs injected into 1-8 cell stage embryos diffuse rapidly to most parts of the embryos, affecting development of the entire organism at a very early stage
(Kim et al., 2010), so the tissue-targeting efficiency is low. The vertebrate retina is derived from the diencephalon of the brain (Moorman, 2001). If early brain development is affected by injection of unmodified MOs, defects in the retina could due to abnormal development of the brain. Therefore, using this technique (i.e. injection of pcdh17 MOs into 1-8 cell stage embryos), it is hard to draw conclusion whether the observed retinal defects in the pcdh17 morphants were caused by the MOs’ direct function in the retinal cell differentiation or indirect function by affecting brain development, which then affects the retinal development. Moreover, the conventional (unmodified) MOs cannot be applied to specific areas in older or adult organism because such large polar structures cannot penetrate the cell membrane (Li and Morcos, 2008). Vivo-MOs are modified MOs with guanidinium headgroups of the arginine residues attached to conventional MOs. The
60 guanidinium headgroups on the vivo-MOs interact with the phosphate or other anionic moieties on the liposomal or cell membrane to allow vivo-MOs get into cells (reviewed by Pantos et al., 2007). Seven to fifteen guanidine headgroups/MO are optimal for efficient cellular uptake (Rothbard et al., 2004; Morcos et al., 2008; Moulton and Yan,
2008). Therefore, vivo- MOs can be applied to study gene functions in specific organs in older embryos, larvae or adult organism. This approach has recently been successfully used to study both skeletal and cardiac muscles in mdx mice in vivo (Moulton and Jiang,
2009; Wu et al., 2009) and in zebrafish (Kim et al., 2010).
To study a more direct role of pcdh17 in zebrafish retinal development, I injected vivo-pcdh17sMO in the right eye of 25-28 hpf embryos, followed by assessing eye development in these embryos. At this stage, retina is completely separated from the brain (the first retinal axons have just begun to be generated), therefore development of the brain unlikely affects eye development. The similar eye (smaller) and retinal cells defects (e.g. reduced RGC layer and thinner optic nerve) observed in these embryos as those injected with the unmodified pcdh17 MOs, suggesting that pcdh17 function directly on zebrafish retinal cell differentiation. The results that the uninjected eye (left eye) of these embryos was also smaller compared to control embryos with no injection to that eye suggest that the vivo-pcdh17sMO must have reached to the other eye likely via blood circulation. Indeed, vivo-MOs have been successfully used to study genes functions in adult organisms by intravenous injections to deliver MOs systematically, and stronger
MOs effects have been obtained from localized injection to specific tissue/organ
(Moulton and Jiang, 2009), which may explain, at least partially, the more severe retinal defects in the injected right eye compared to the uninjected left eye.
61 Possible mechanisms of pcdh17 function in retinal development
It is possible that pcdh17 function in retinal cell development through controlling expression of regulatory molecules known to be involved in the vertebrate retinal differentiation. Expression of crx, neuroD and otx5 in the morphant retina is reduced compared to control retina, which is similar to results obtained from embryos with disrupted cdh2, cdh4, or cdh6 function (Babb et al., 2005; Liu et al., 2007; 2008). Also similar to cdh2 glo mutants (Liu et al., 2007), crx expression is especially reduced in the posterior half of the pcdh17 morphant retina. But otx5 expression in the glo mutant is almost absent in the posterior half of the retina, whereas otx5 expression is only slightly reduced compared to the anterior portion of the retina in the pcdh17 morphant. Like cdh6 morphants (24 hpf, Liu et al., 2008), rx1 expression was unchanged in the 24 hpf pcdh17 morphant retina. But unlike the cdh6 morphant retina where neuroD and crx expression is barely detectable, while otx5 expression is mainly confined to the anterior ¼ of the retina
(Liu et al., 2008), expression of these molecules is only moderately affected in the pcdh17 morphant retina. In summary, reducing cadherins functions diminish expression of most of these transcription factors, but each cadherin has differential effect on expression of these regulatory molecules. As mentioned above, the retinal defects are more severe in cdh6 morhpants than pcdh17 morphants. Not surprising, there seems to be a correlation between extent of reduction in the expression of these molecules (excluding rx1) and severity in the retinal defects.
Pathway analysis is important for explaining mechanisms underlying biological phenomenon and gene functions. My proteomics data showed that pcdh17 morphants exhibited lower metabolic levels, less molecular phosphorylation activities and more 62 protein degradation compared to control embryos. One of my main proteomics findings is that the pcdh17 morphants had greatly reduced GFAP expression compared to control embryos, which was further confirmed using ICC on tissue sections.
GFAP is the major protein of glial intermediate filaments expressed by differentiated Müllar glia (Danhl, 1976; Marcus and Easter, 1995). It has been used as a marker for brain and retina radial glial development (Alunni et al., 2005). Previous researches have shown that Müller glia function as stem cells in the retina (Bringmann et al., 2006). In zebrafish retina, differentiated Müller glia cells can enter the cell cycle and produce retinal progenitors that differentiate as either rod or cone photoreceptors
(Bernardos et al., 2007). Failure in its development can cause abnormal retinal neuron differentiation and causes loss of vision. Interestingly, the morphant retina showed more reduction in GAFP staining compared to control retina, while there was no detectable difference in GAFP expression in the hindbrain between the morphants and control embryos. However, since proteomics showed more than two- fold reduction in GFAP in the morphants, it’s unlikely that the reduction in the eye would be sufficient to account for this much reduction in the whole embryos. Besides the retina, GFAP is also expressed in other regions of the body, including the forebrain and spinal cord. Therefore GFAP expression is also likely to be reduced in other GFAP containing regions (except the hindbrain) in the morphants.
In addition, presence of GFAP is associated with increased expression of some extracellular matrix and adhesion molecules (Abd-el- Basset et al., 1992). Pcdh17 may control the early axon outgrowth and cell differentiation through changing the adhesive interaction between its extracelluar domain and extracellular matrix. Decreased GFAP
63 expression may result in less adhesion between pcdh17 and extracellular matrix or other adhesion molecules.
Many classical cadherins have been shown to control cell motility, growth and differentiation through β-catenin/TCF/LEF signaling pathway (Peifer et al., 1992;
Clevers, 2006; Delva and Kowalczyk, 2009). Protocadherins (e.g. pcdh24) were shown to regulate cell activities through the same pathway (Ose et al., 2009). Beta-catenin expression appears to be reduced in pcdh17 morphant retina (Fig. 9). Actin-like protein, another functional protein in cell motility and signaling, was also down-regulated in the morphant (Table 7, SB132623 and SB132625). This suggests that function of pcdh17 may also follow the β-catenin/TCF/lef signaling pathway.
Since pcdh-17 is a non-clustered pcdh in the δ-pcdh subfamily, it is not clear if it controls cell adhesion and signaling using similar mechanisms as clustered pcdhs such as
γ-pcdhs (Han et al. 2010). Chen and Gumbiner (2006) showed that pcdhs could modify cell adhesion by regulating the adhesion activity of a classical cadherin. More recently,
Jontes’ laboratory showed that pcdh19 interacts with cdh2 to regulate brain development
(Biwas et al. 2010). Therefore, this could be another potential mechanism by which pcdhs affect cell adhesion and signal transduction. Classical cadherins were not among those proteins identified in my proteomics. This may partially due to limitations of the technique, since classical cadherins have large molecular weight (>100 kDa), and 2D gels are not efficient in separating proteins with large molecular weight.
Proteomics has been considered as one of the high-throughput sequencing and gene/protein profile approaches in this area (Graves and Haystead, 2002). However, challenges still exist although the techniques have been developed for decades (reviewed
64 by Khatri et al., 2012). For example, some of my identified proteins only had predicted profile in online databases; some of them have no annotations; even those annotated proteins, they only had general overlapped concepts but no specific targets.
I chose to focus more on analyzing protein expression profiles of 72 hpf embryos, instead of 50 hpf embryos, because of biased representation of various isoforms of vitellogenin, one of the major yolk proteins, in 50 hpf embryos, which were shown to obscure 2D gel images and analysis (Link et al., 2006). The older embryos (72 hpf) have much smaller yolk than 50 hpf embryos, but vitellogenin was still one of the major proteins present in 72 hpf embryos. To minimize yolk protein interference, removal of the yolk (more practical, Link et al., 2006) or use only eye tissue (less practical because of smaller eye size) for proteomics maybe used in future studies.
The information gained from my proteomic study is limited. It is impossible to analyze each differentially stained spot because of high cost. Even with plenty of spots analyzed, challenges still remain as discussed by Kharti et al. (2012). In my analysis, pathways were independently considered, without crossing over and/or overlap. However, in real biological systems, dynamic protein expression and function at different developmental stages and/or under different physiological conditions, often require participation of multiple interacting signaling pathways. Many of the identified differentially expressed proteins in my study belong to metabolic pathways rather than signaling pathways. Furthermore, some of the sequences are still not annotated. Some of them are predicted (histone H4-like in SB132628, Table 7; profilin-1 isoform 1 in
SB13356, Table 7). Incomplete annotations, low quality and inaccurate existing annotations are some of the main problems when performing databases searching.
65 We have begun to use microarray, another widely used but more high-throughout approach, to study differential gene expression between control embryos and pcdh17 morphants. It will be interesting to match results from my proteomics with that from microarray to see if similar molecules and pathways are identified by both methodologies.
This combined approach will likely help us better understand how pcdh17 function at the molecular level in regulating zebrafish eye development. The preliminary results from the microarray showed down regulation of classical cadherins, clusterer protocadherins and weak homophilic adhesive molecules such as tyrosin kinase Fyn, alpha isoform of protein phosphatase 1, or adaptor protein Disabled 1 in the morphant group, indicating that pcdh17 interacts with these molecules to regulat zebrafish retinal development (Frank and Kemler, 2002; Nakao et al., 2008; Ose et al., 2009). Further analysis of the microarray data using Gene Ontology and other pathway analysis software should provide us with more information on possible molecular mechanisms underlying pcdh17 function in the differentiation of the retinal cells.
Summary and perspective
In this study I showed that pcdh17 is expressed during critical stages of zebrafish retinal development. Blockage of pcdh17 function using either unmodified or modified
(i.e. vivo-) zebrafish pcdh17 specific MOs resulted in greatly reduced eye size (due to reduced retinal cell proliferation), disrupted differentiation of RGCs, amacrine cells and photoreceptors. Reduced expression of selective retinal regulatory molecules in the pcdh17 morphants suggests that pcdh17 may control retinal development through affecting expression of these regulatory molecules. Molecular mechanisms underlying
66 pcdh17 function in the retina was further studied using proteomics, which identified several proteins, including GFAP, had altered expression in the pcdh17 morphants. It will be interesting to find if results from our current microarray analysis of pcdh17 morphants would confirm my proteomics findings, and identify more molecules whose expression are greatly changed when pcdh17 expression is reduced, providing more information on possible mechanisms underlying pcdh17 function in zebrafish retinal development.
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