Initial Molecular Characterization of Planarian Pigment Cells, a New Model for Studying Lineage Specification and Gut Excretion by Xinwen He A thesis submitted in conformity with the requirements for the degree of Master of Science Department of Molecular Genetics University of Toronto © Copyright by Xinwen He 2016

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Initial Molecular Characterization of Planarian Pigment Cells, a New Model for Studying Lineage Specification and

Gut Excretion


Xinwen He

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Molecular Genetics University of Toronto

© Copyright by Xinwen He 2016


Initial Molecular Characterization of Planarian Pigment Cells, a

New Model for Studying Lineage Specification and Gut Excretion

Xinwen He

Master of Science

Department of Molecular Genetics

University of Toronto



ASCs play crucial roles in tissue homeostasis and regeneration. Here, I established pigment cells

in Schmidtea mediterranea as a novel model to study ASC lineage specification by identification

of hmbs and alas as makers for pigment cells, and ans as a marker for their progenitors. Co-

expression analysis between hmbs and tryptophan metabolism enzyme, kmo, suggests

ommochrome as one of the responsible pigments for planarian body pigmentation. Initial

characterization of this lineage reveals an fst homolog as a negative regulator of pigment cell

regeneration and proliferation. Functional studies of light-induced depigmentation also uncover

homologs of draper and epimerase as positive regulators of gut excretion mediated pigment cell

removal. Together, these findings in pigment cell lineage and gut excretion pave ways to future

studies of ASC regulations.



Life is about passing down messages. Primitive organisms pass down their messages in

the form of genetic materials. We human beings pass down our messages to our future

generations, not only with our genetic materials, but also in the forms of knowledge and

experiences. This thesis presented here is a recording of the knowledge and experiences I

gathered through my shallow studies in planarians, through the guidance from my supervisor,

and through working and living with my colleagues, families, and friends.

First and foremost, I would like to thank my supervisor Dr. Bret Pearson for his

continued support and guidance through my scientific endeavors. Using his own knowledge and

experiences, he showed me how to design and perform experiments, and how to think critically

and scientifically. More importantly, he is a great role model that inspires me on the meanings of

my career and personal life. It is my privilege to work with him, to do science together, and to

make discoveries that will benefit our society and the future generations. I would also like to

thank my supervisory committee, Dr. Marc Meneghini, and Dr. Nadeem Moghal, as well as my

collaborator, Dr. Jason Pellettieri. I have benefited greatly with their scientific expertise in their

fields and their different perspectives on the project. I am grateful to all past and present

members of the Pearson Lab. My work is only possible because of their optimization of

techniques, their advice on scientific problems, and the enjoyable lab environment they created.

I am sincerely thankful for my parents and grandparents. Their constant support since my

childhood has accompanied me to travel around the globe. I can never forget the encyclopedias

they bought me when I was little, which opened the doors for me to the magnificent world of

science. I am extremely lucky to have such a great family and have the resources and support for

me to freely develop my career and become the person I want to be. I would also like to thank

my girlfriend, Kelly. She is the best friend that I have ever had in my life. For over eight years,

her constant motivation, support, and love are the key to helping me staying focused on my


Last but not least, I thank all the planarians that have sacrificed for this study. Their lives

perished with a meaning. And I hope the messages of the knowledge generated can be passed on.


Table of Contents

Acknowledgments.......................................................................................................................... iii

Table of Contents ........................................................................................................................... iv

List of Figures ................................................................................................................................ vi

List of Appendices ........................................................................................................................ vii

Introduction .................................................................................................................................1

1.1 ASCs are sources for tissue homeostasis and regeneration .................................................1

1.2 Freshwater planarian, Schmidtea mediterranea, boasts distinct properties as a suitable

model for ASC and regeneration study ................................................................................4

1.3 Advances and drawbacks of current approaches in planarian based ASC and

regeneration studies .............................................................................................................5

1.4 Ablation of planarian pigment cells by light exposure allows dissection of specific

lineage specification regulators ............................................................................................6

1.5 Previous studies on planarian pigment cells are limited and contradictory .........................6

1.6 Gut excretion of various cell types in response to RNAi and light exposure treatment ......8

1.7 Outline and rationale for the thesis research ........................................................................9

Materials and Methods ..............................................................................................................10

2.1 Animal husbandry and light-induced depigmentation .......................................................10

2.2 Intact planarian sample collection and RNA deep sequencing ..........................................10

2.3 Candidate selection and cloning for expression and functional analysis ...........................11

2.4 Riboprobe synthesis for expression analysis .....................................................................12

2.5 WISH and dFISH for expression analysis .........................................................................13

2.6 Targeted functional knockdown by RNA interference ......................................................16

Results .......................................................................................................................................17

3.1 Identified hmbs and alas as planarian pigment cell markers .............................................17

3.1.1 Selection of candidates and their expression patterns............................................17

3.1.2 Genes with sub-epidermal expressions and pigment cell-like morphologies ........19


3.1.3 Expression of spider markers correlates to pigment status of planarian tissue ......20

3.1.4 hmbs is functionally required for pigment production and maintenance ...............22

3.1.5 hmbs is co-expressed with other spider markers and kmo .....................................23

3.1.6 Pigment cells show inward migration toward gut during depigmentation in

response to light exposure ......................................................................................26

3.1.7 Pigment cells display fast regeneration timeline during repigmentation ...............28

3.1.8 Decreasing levels of co-expression between punctate and spider markers as

repigmentation progresses .....................................................................................30

3.2 Identified regulators of depigmentation and pigment cell regeneration ............................32

3.2.1 hmbs is not required for pigment cell survival .......................................................32

3.2.2 Presence of pigment is required for light-induced pigment cell loss .....................32

3.2.3 epimerase positively regulates depigmentation in response to light .....................34

3.2.4 draper positively regulates depigmentation in response to light ...........................34

3.2.5 Disruption of draper function leads to formation of wrinkles on epidermis and

ectopic expressions of lineage markers ..................................................................36

3.2.6 follistatin negatively regulates pigment cell regeneration and homeostasis ..........37

Discussion .................................................................................................................................40

4.1 Identified markers, and established pigment cells as a novel model for ASC and

regeneration study ..............................................................................................................40

4.2 Regulation of depigmentation and pigment cell regeneration ...........................................43

4.3 Future directions ................................................................................................................46

4.3.1 Deeper analysis of pigment cell and potential progenitors ....................................46

4.3.2 Continued screening of pigment cell lineage markers and regulators ...................46

4.3.3 Further analysis of depigmentation regulators .......................................................48

4.3.4 Further analysis of pigment cell regeneration regulators .......................................48

4.3.5 Comparative transcriptomics for identification of gut excretion regulators ..........49

References ......................................................................................................................................50

Appendices .....................................................................................................................................57


List of Figures

Figure 1 Decision making of adult stem cells. ............................................................................ 2

Figure 2 Planarian pigment cell as a model of lineage specification and gut excretion. ............ 7

Figure 3 RNAseq sample collection. ........................................................................................ 11

Figure 4 Expression patterns of genes with differential expression levels in depigmented

animals. ...................................................................................................................... 18

Figure 5 Expression patterns of spider and punctate genes resemble pigment cell morphology.

.................................................................................................................................... 20

Figure 6 Expression of spider and punctate genes directly correlates to the pigment status of

planarian tissue. .......................................................................................................... 21

Figure 7 Functional requirement of hmbs in pigmentation maintenance and repigmentation. 23

Figure 8 Co-expression analysis of spider and punctate markers. ............................................ 25

Figure 9 Pigment cells migrate towards gut during depigmentation. ....................................... 27

Figure 10 Rapid regeneration of pigment cells during repigmentation. ..................................... 29

Figure 11 Co-expression of spider and punctate genes decreases along repigmentation time

course. ........................................................................................................................ 31

Figure 12 hmbs is not required for pigment cell survival and its RNAi offers protection from

light exposure. ............................................................................................................ 33

Figure 13 RNAi phenotypes of epimerase, draper and fst. ........................................................ 35

Figure 14 Ectopic expression of lineage markers in draper RNAi condition. ............................ 38

Figure 15 Model for pigment cell lineage proliferation.............................................................. 42

Figure 16 Model of epimerase, draper and fst functions in pigment cell lineage specification

and excretion. ............................................................................................................. 45


List of Appendices

Appendix 1 List of in situ patterns identified but not included in the main text. ..................... 57

Appendix 2 List of genes cloned and screened in this study. ................................................... 58

Appendix 3 List of phenotypes observed during RNAi screen. ............................................... 61



1.1 ASCs are sources for tissue homeostasis and regeneration

Adult stem cells (ASCs) are the ultimate source for the tissue turnover of approximately 50 – 70

billion cells per day in humans (Reed, 1999). Many mysteries lie behind the phenomenal power

of ASCs, not only can they regulate the exact type, and the exact number of differentiated cells

to replace, but also can they maintain a stable population of themselves in the long term.

Terminal selector transcription factors play key roles in determining the types of differentiated

cells. In some cases, a single terminal selector can regulate hundreds of downstream effector

genes, and its ectopic expression can even reprogram cells to adopt a different fate (Eastman et

al., 1999; Flames and Hobert, 2009; Hobert, 2011). For instance, unc-30, a C. elegans homeobox

gene, controls the expression of crucial genes, including unc-25 and unc-47, for the identity and

function of GABAergic neurons (Eastman et al., 1999). And the ectopic expression of unc-30 is

capable of driving wild-type unc-25 and unc-47 promoters in other tissue types (Eastman et al.,

1999). Despite a relatively rich knowledge of terminal selector transcription factors for the

differentiation of neuron types, understanding of such mechanisms in many other tissue types is

lacking. In addition to the determination of tissue types, what is even more poorly understood is

the regulation of the exact amount of differentiated cells to replace. The feedback control

between differentiated cells and ASCs is one interesting mechanism to be investigated, a

mechanism that could be compromised in the case of cancer development.

In order to maintain a stable population of ASCs, establishing an asymmetry between the

proliferation and differentiation is crucial (Figure 1) (Sherley, 2002; Simons and Clevers, 2011).

Such asymmetry can be achieved intrinsically, by disproportionate distribution of RNA and

protein molecules that drive different cell fates (Bossing et al., 1996; Doe, 1996, 2008; Doe and

Bowerman, 2001; Bayraktar, 2010). For example, Inscuteable, Prospero, and Numb are

asymmetrically localized to the apical cortex in neuroectodermal cells in Drosophila embryos

and physically segregated to daughter cells after neuroblast divisions (Doe, 1996). Prospero,

with a homeodomain and nuclear localization signals, then drives the differentiation and

suppresses the stem cell identity in the daughter cells (Doe, 1996; Bayraktar, 2010). Asymmetry

can also be achieved extrinsically, such as receiving different signals from the stem cell and

differentiated tissue microenvironment (Snippert et al., 2010; Sato et al., 2011). For example, the


Figure 1 Decision making of adult stem cells. Adult stem cells (ASCs) respond to clues such as aging and injury, then decide between symmetric and asymmetric proliferation to fulfill the different functions during homeostasis and regeneration. ASCs also receive feedbacks from daughter cells to control the rate and mode of proliferation.

Lgr5 stem cell fate is determined by signaling in the microenvironment created by the

differentiated Paneth cells in the mammalian intestinal crypt (Sato et al., 2011).

Adding to the complexity of the asymmetry of ASCs is that a re-balance between proliferation

and differentiation is necessary for the survival and well-being of an organism in response to

injury, aging, and disease. Regeneration, an ability that we humans desired and has captured our

imagination since Greek mythology, is a fascinating phenomenon to showcase the shift of

balance between the different fate choices of ASCs. Regeneration is also a very poorly

understood question, given its huge potential to shed light on tissue differentiation, remodeling,

and the control of tissue size and proportions. One of the first scientific observations of

regeneration is made in Hydra by Abraham Trembley in 1740. The simple organism was

observed to completely regenerate its head after amputation (Lenhoff and Lenhoff, 1986). Soon


after, discoveries of regeneration in more and more complicated organisms were made, including

planarians, amphibian tadpoles, and salamanders (Pallas, 1766; Spallanzani, 1769; Sánchez

Alvarado A, 2000). And more than 200 years later since Trembley’s initial discovery, the

regeneration ability of human liver was noticed (Widmann and Fahimi, 1975). Many different

types of regeneration exist and can be categorized into two categories based on their requirement

of blastema formation. The blastema is a mass of morphologically undifferentiated cells formed

at the end of the stump during regeneration, similar in form and function to the vertebrate

embryonic limb bud (Sánchez Alvarado A, 2000; Tasaki et al., 2011). For instance, human liver

regeneration requires limited proliferation of the surviving cells while bone regeneration requires

proliferation and differentiation of stem cells already present in the tissue (Michalopoulos and

DeFrances, 1997; Prockop, 1997). Both types of regeneration are non-blastema based. In

comparison, more robust regeneration programs involve blastema formation (Sánchez Alvarado

A, 2000). Examples of this type of regeneration are used by planarians, mollusks, and also by

vertebrates during limb and tail regeneration (Brøndsted, 1969; Newth, 1958; Needham, 1952;

Lange, 1920; Brockes, 1997; Iten and Bryant, 1976; Morgan, 1901). Previous studies have

established the developmental programs across the animal kingdom as a conserved mechanism

(Wolpert, 1994; De Robertis and Sasai, 1996). It has also been shown that the regeneration

program shares a lot of similarities between the tissue differentiation and proliferation control

during embryonic development (Needham, 1952). It is very likely that the regeneration program

is also a conserved mechanism and has common origins to a wide variety of regeneration

phenomena. Thus, studies of regeneration in different model animals can be cross-related.

The understanding of ASC and regeneration biology would not only allow us to gain insights to

the elegant developmental and differentiation programs of multicellular organisms but also to

better appreciate many disease states and realize the therapeutic potentials in regenerative

medicine. Despite these great implications, many areas of these subjects await further

investigations. It is of my interests in this thesis research presented here to gain more insights on

a few key steps toward a better understanding of the ASC biology and regeneration. Specifically,

by identifying and investigating the regulators of ASC asymmetric proliferation and

differentiation, regulators that determining the fate commitment during lineage specifications,

and the mechanisms mediating the feedback control between differentiated tissue and ASCs

during homeostasis and during regeneration (Figure 1).


1.2 Freshwater planarian, Schmidtea mediterranea, boasts distinct properties as a suitable model for ASC and regeneration study

ASCs are rare and hard to access in almost all model organisms, making in vivo studies of ASC

biology extremely difficult. Common model organisms, including Drosophila melanogaster,

Caenorhabditis elegans, Mus musculus, lack robust regeneration events to be studied. While

animals traditionally used to study regeneration, including axolotls and salamanders, are

extremely refractory to genetic and molecular manipulations (Sánchez Alvarado A, 2000). In

comparison, the freshwater planarian, Schmidtea mediterranea, hosts a large and accessible

population of ASCs (neoblasts), representing 10-20% of their cells, has robust regeneration

programs, and recent advances in molecular biology techniques in these animals really made

them an ideal model for ASC and regeneration studies (Morgan, 1901; Krichinskaya and

Martynova, 1975; Hayashi et al., 2006; Wagner et al., 2011; Saló and Baguñà, 2002; Sánchez

Alvarado A. 2003, 2006; Cebrià, 2007; Rossi et al., 2008; Saló et al., 2009; Aboobaker, 2011;

Elliott and Sánchez Alvarado, 2013; Rink, 2013). S. mediterranea is a type of free-living

flatworm belonging to the Lophotrochozoan superphylum (Aguinaldo et al., 1997). Standing at

this unique phylogenetic location in relation to mammals while hosting a wide variety of

homologs of essential genes to humans, planarians are an ideal model for studies to provide us

with valuable information in the conservation and evolution of ASCs and regeneration regulators

(Labbé et al., 2012). Showcasing their large population of ASCs, planarians display amazing

regeneration capabilities, with reported regeneration into an entire functional animal from an

amputated piece of as little as 1/279th of the body mass of the original animal (Morgan, 1901).

Regeneration of an entire planarian body is extraordinary from limited ASCs in small amputation

fragments, especially when one considers the extremely complicated anatomy of the flatworm,

with a central brain and a PNS that contain all neuron types that can be found in an invertebrate

system, a muscular pharynx and a triclad gut for uptake of food and nutrient intake involving

phagocytosis, and a primitive excretory system protonephridia (Cebrià, 2007; Cebrià et al., 2002;

Reddien and Sánchez Alvarado, 2004; Forsthoefel et al., 2011; Scimone et al., 2011). In addition

to the robust regeneration, planarians are also able to replace their aging tissue with a fast

turnover rate leading to an immortal lifestyle (Eisenhoffer et al., 2008; Pellettieri and Sánchez

Alvarado, 2007). How planarians balance their regeneration/immortality without tumor

development is an interesting question as well (Pearson and Sánchez Alvarado, 2008). Finally, S.


mediterranea is a constitutive adult, thus allowing a number of gene functions to be investigated

that may be required during embryonic development in other model organisms (Newmark and

Sánchez Alvarado, 2002). Combining all the above attributes, planarians are an ideal model for

studying the molecular mechanisms underlying regeneration, and ASC proliferation and


1.3 Advances and drawbacks of current approaches in planarian based ASC and regeneration studies

Research in S. mediterranea has greatly expanded in the past decade as a result of the increasing

availability of molecular and genetic tools. Available techniques include whole-mount in situ

hybridization (WISH) to investigate gene expression patterns and double fluorescent WISH

(dFISH) to perform co-localization study on gene products (Pearson et al., 2009). Planarians are

also amenable to gene function study by RNA interference (RNAi), where double strand RNA

(dsRNA)-expressing bacteria are fed to the animals and subsequently spreads to all tissue types

(Sánchez Alvarado and Newmark, 1999). The S. mediterranea genome has been sequenced and

five transcriptomes of this species have been generated (Robb et al., 2008; Robb et al., 2015;

Labbé et al., 2012; Zhu et al., 2015; Solana et al., 2012). These sequencing datasets made

possible to perform a number of comparative transcriptomic studies to dissect the involvement of

different genes under different conditions. All differentiated planarian tissue are post-mitotic and

the ASCs are the only dividing cells in the animal, allowing researchers to use gamma irradiation

to specifically ablate all ASC population (Eisenhoffer et al., 2008). A number of studies have

been published using this technique to analyze the gene expression in ASCs versus differentiated

tissue (Eisenhoffer et al., 2008; Zhu et al., 2015; Reddien et al., 2005). Another technique is to

induce planarian regeneration, most commonly by amputation, to observe changes in gene

expression in wound response. A number of studies were able to identify regulators of ASC

proliferation and migration, and of program cell death during tissue remodeling (Kao et al.,

2013; Pellettieri et al., 2010; Gaviño et al., 2013; Currie and Pearson, 2013). The amputation

method, however, causes loss of multiple tissue types, making it difficult to dissect the

involvement of regulators in individual lineage specification. Methods that allow identification

for ASC regulators following a specific cell type loss, without inducing physical injuries, are

urgently needed.


1.4 Ablation of planarian pigment cells by light exposure allows dissection of specific lineage specification regulators

S. mediterranea displays a brownish color under normal laboratory conditions (Figure 2A)

(Newmark and Sánchez Alvarado, 2002). It was accidentally noted that planarians left under

light exposure become depigmented over the course of several days (Figure 2A). It is also known

that planarians display a strong negative phototaxis and prefer a dark environment. It was further

observed that increased level of apoptosis surrounds the gut area of the animals while the gut was

darkened at the same time, presumably enriched in excreted pigment or pigment cells (Figure

2B, C. Apoptosis assay by terminal deoxynucleotidyl transferase dUTP nick end labeling

(TUNEL), courtesy of Dr. Jason Pellettieri). When depigmented planarians are removed from the

acute light exposure, however, the animals will repigment over the course of approximately two

weeks (Figure 2A). Thus, I hypothesize that planarian pigment cells are lost in response to acute

light exposure and that the repigmentation following acute light exposure requires rapid

regeneration of pigment cells. Thus, the planarian light-induced depigmentation provides us with

a possible exciting model to investigate the ASC regulation in response to a specific type of

tissue loss, the pigment cell loss, without physical injury, allowing us to identify regulators of

this specific lineage.

1.5 Previous studies on planarian pigment cells are limited and contradictory

Surprisingly little is known about planarian pigment cells, let alone regulators of their lineage

specification. Previous studies came to different conclusions about the composition of planarian

pigment. One study in a related planarian species, Dugesia gonocephala s. l, show that planarian

pigment cells have process-rich morphology, similar to that of melanocyte in the mammalian

epidermis (Palladini et al., 1979a). Enriched in these processes that extend to the sub-epidermal

layer between muscle fibers are pigment granules (Palladini et al., 1979a). Each pigment cell is

thus responsible for the pigmentation of a much larger surface area of the animal than that of the

pigment cell soma alone. Interestingly, melanocyte-stimulating hormone release-inhibiting factor

(MIF) was shown to induce a lighter pigmentation of the treated planarians (Palladini et al.,

1979b). These results suggest melanin to be the potential pigment of planarians. On the other

hand, one study biochemically determined the body pigment of another related planarian species,

Dugesia ryukyuensis, to be ommochrome, with the major evidence coming from the absorption


Figure 2 Planarian pigment cell as a model of lineage specification and gut excretion. A) Planarians under normal laboratory conditions exhibit a brownish body pigmentation. After ~10 days of light exposure, planarians are depigmented. The body pigmentation recovers in ~2 weeks following removal of light exposure. B) Planarian gut darkens during depigmentation (lower left) and an increased level of apoptosis was detected by TUNEL (upper right). The increased level of TUNEL staining is associated with the darkened gut (upper left). C) Details of TUNEL stains overlapping with darkened gut. D) Outline of major goals for the thesis study. 1. Establishing molecular markers for planarian pigment cells and their progenitors. 2. Identifying regulators of pigment cell differentiation, regeneration and gut excretion. (Live imaging and TUNEL stain images (Panel B and C) courtesy of Dr. Jason Pellettieri.)


maxima of the extracted body pigment at 256nm, 367nm, and 463nm, characteristic of

ommochromes (Hase et al., 2006). We know from studies of pigment in other organisms that

melanin and ommochromes are derivatives of tyrosine and tryptophan metabolism, respectively

(Ando et al., 2007; Croucher et al., 2013). It is of my interest to investigate perturbation in which

of the two amino acid metabolism pathways would affect planarian pigmentation.

1.6 Gut excretion of various cell types in response to RNAi and light exposure treatment

Planarians are acoelomates without body cavities, neither do they have specialized circulatory or

respiratory systems (Carranza et al., 1997). These attributes limit planarians to adopt a flattened

body shape for diffusion of oxygen and nutrients. Their triclad gut, although having only one

opening for both ingestion and egestion, is one of the few specialized organs that cover the entire

length and width of the animals (Forsthoefel et al., 2011). One would speculate if planarian gut

plays important roles that are fulfilled by the circulatory or lymphatic system in other animals.

Indeed, planarian gut has been noted to take part in the excretion of cells under various

conditions, a typical role played by lymphatic system in vertebrates. Under p53 RNAi condition,

planarians display an over-proliferation of stem cells, while at the same time, an ectopic

expression of cyclinB positive cells can be observed in the gut cavity of the animals (Pearson and

Sánchez Alvarado, 2010). It is unknown whether the cell excretion function of planarian gut is

one of the mechanisms used to remove detrimental cells and guard against tumor development.

In the same study, when p53 RNAi was applied to animals following amputation, head and tail

fragments, which no longer possess a pharynx and thus no opening of the gut to the external

environment, were noticed to develop tissue dysplasia (Pearson and Sánchez Alvarado, 2010).

No such tissue dysplasia was observed in the trunk fragment, however, presumably because the

gut is still open due to the presence of the pharynx on these fragments and able to remove the

detrimental cells excreted by the animal. Another event of gut excretion of cells was noticed

under mex-3 RNAi condition, which causes loss of progeny cells and eventually the failure of

tissue turnover (Zhu et al., 2015). Progeny cells marked by prog-2 in the parenchyma were

ectopically found inside the gut and eventually excreted (Zhu et al., 2015). Considering these

previous observations, the darkening and increased apoptosis activity in planarian gut during

acute light exposure is likely to be gut excretion as well, and it is of my interest to identify


regulators of this process. In comparison to previous RNAi-based approaches, the light exposure

induced depigmentation offers a novel model on gut excretion.

1.7 Outline and rationale for the thesis research

In collaboration with Dr. Jason Pellettieri’s lab (Keene State University, NH, USA), we have

generated an RNA-deep sequencing (RNAseq) dataset of S. mediterranea, by collecting intact

animals at several time points during the acute light exposure induced depigmentation and the

repigmentation process 4 days after returning to the dark. Comparing the RNAseq results

between the Day 10 (completely depigmented) time point and the control, expression levels of

more than one hundred transcripts were either increased or decreased by more than two-folds.

Working with these candidates of differentially-expressed transcripts, I have two goals in this

thesis research. First, I will establish the first molecular markers for planarian pigment cells

(Figure 2D). I hypothesize that a number of transcripts with decreased expression level in the

depigmented animals are specifically expressed in the pigment cells and can be demonstrated to

be pigment cell markers. My rationale is that if pigment cells are lost during light exposure, the

specifically expressed transcripts are no longer present in the animals, thus showing a decreased

expression level in the RNAseq data of depigmented worms. Second, I aim to identify regulators

of planarian pigment cell lineage specification, regeneration, and gut excretion (Figure 2D). I

hypothesize that certain genes with altered expressions, either with increased or decreased

expression in planarians in response to light exposure are key regulators of these processes. My

rationale is that planarians would activate specific genetic machinery to replenish pigment cells

after sensing a reduction in the pigment cell population and that the active removal of pigment

cells through gut excretion is mediated by genes that are specifically regulated during light

exposure. Both of the two processes would lead to altered expressions of responsible genes that

can be identified in the RNAseq data. To the end of these two goals, I performed a screen of 40

up-regulated and 69 down-regulated genes in the depigmented animals. In addition, I also

screened a few candidate genes with suspected involvement, by analyzing their expression

patterns and their functional involvement during depigmentation, gut excretion, pigment cell

lineage specification and regeneration.


Materials and Methods

2.1 Animal husbandry and light-induced depigmentation

A diploid, asexual CIW4 strain of Schmidtea mediterranea originally collected in Barcelona,

Spain was reared as previously described (Benazzi et al., 1972; Newmark and Sánchez Alvarado,

2002). The animals used in this study were kept in a dark incubator at 20°C under default/control

conditions. The animals were cultured in planarian water (Milli-Q H2O supplemented with 21g/L

Instant OceanTM, 0.9mM NaHCO3, 0.83mM MgSO4, 0.9mM CaCl2 and 0.04mM KHCO3 salts,

as previously described; Zhu et al., 2015) Animals were fed with beef liver paste every 10 days,

but a starvation period of one week was carried out before performing experiments on the animal

subjects. For the experiment of collecting whole planarian samples for RNAseq during the light-

induced depigmentation and repigmentation time course, a light exposure set up in a Styrofoam

box using CFL bulb was used. Specifically, a 14W, 2700K CFL bulb was placed approximately

30cm away from a tissue culturing plate containing the planarians. A piece of UV-resistant glass

was placed between the tissue culturing plate and the light source. A fan was used in this set up

to keep the circulation of the air and to prevent overheating of planarians. The animals were

treated with 5 cycles of 24 hours of light exposure and 24 hours of dark exposure (dark exposure

by placing the samples in the normal condition incubator) for a total of 10 days, at which point

the animals were completely depigmented on their dorsal surface. The animals were then

returned to the normal condition dark incubator for the rest of the experiment and to be observed

for repigment. For the light-induced depigmentation performed during the RNAi functional

screens, an alternative LED light exposure set up was used. The LED set up bares the sample

principle of the CFL set up, and unless otherwise noted, a cycle of 15-minute light and 225-

minute dark exposure was performed for 48 times (a total of 8 days at 6 cycles per day). This

alternative setup and light exposure schedule were used in the RNAi screen instead for its

shortened timeline and more consistent animal health at the end of the treatment.

2.2 Intact planarian sample collection and RNA deep sequencing

In collaboration with Dr. Jason Pellettieri’s Laboratory, 5 to 10 intact planarian animals were

collected from 3 biological replicates under the CFL light-induced depigmentation set up for

total RNA extraction using TRIzol® Reagent (Invitrogen, catalog number 15596-026) (Figure 3).

Two time points during the depigmentation, at Day 1 (24hr) and Day 2 (48hr), one time point


when the worms are completely depigmented, at Day 10, one time point when the worms are

repigmenting, at Day 14 (since start of light exposure treatment, or Day 4 repigmentation), in

addition to a control sample with no light exposure, for a total of five samples were collected

(Figure 2). Poly-A-selected cDNA libraries were prepared from the purified total RNA samples

using TruSeq kits from Illumina. We generated approximately 108 of 50-base single end reads

for each time point sample on Illumina HiSeq2500 with v4 chemistry. Using bowtie2 alignment,

the RNAseq reads were mapped to the 2011 Pearson Lab asexual Schmidtea mediterranea

transcriptome and generated a coverage of approximately 8 times for every nucleotide of the

82,188 transcripts. Pseudocounts of +1 were added to every numerator and denominator when

calculating expression ratios from the RNAseq counts (i.e. raw counts per kilo million (RPKM)

+ 1 was used), in order to reduce the bias of lowly expressed transcripts having large fold

changes without substantial absolute changes.

Figure 3 RNAseq sample collection. Animals were treated with 5 cycles of 24 hours of light and 24 hours of dark exposure for a total of 10 days. After 10 days, animals were placed in the dark for recovery of pigmentation. Whole worm samples were collected at 24 hrs, 48 hrs, 10 days time points during depigmentation, at 4 days time point during repigmentation and at 10 days time point for control animals. Total RNA was extracted and then subjected to cDNA library preparation and RNA deep sequencing.

2.3 Candidate selection and cloning for expression and functional analysis

Candidate genes to be screened for pigment cell markers and regulators of pigment cell lineage

specification were selected using a 2-fold expression change (RPKM + 1) cut-off, from genes


with either increased or decreased expression in Day 10 (depigmented) animals in comparison to

control animals. Near full-length transcript of the established candidate genes were cloned using

gene-specific forward and reverse primers into a double stranded RNA expression vector (pT4P)

as previously described (Rink et al., 2009). For transcripts longer than 1.5kb, up to 1.5kb of 5’

sequences of the transcripts were cloned. Forward and reverse primers were designed to have an

18-22 nucleotide identity to the binding region of the transcript with a melting temperature

around 54°C, then additional sequences of 5’-CATTACCATCCCG-3’ and

5’-CCAATTCTACCCG-3’ were added to the 5’ of forward and reverse primers, respectively.

Total RNA was extracted from 7-day planarian regeneration time course and used as the

template to generate cDNA by SuperScript III Reverse Transcriptase (Thermo Scientific, Catalog

number: 18080-051). Polymerase chain reaction (PCR) was then performed with the forward and

reverse primer sets using the cDNA as the template. PCR products were gel extracted and treated

by T4 DNA polymerase (Thermo Scientific, Catalog number: EP0061) with dCTP as the only

nucleotide available in the reaction mix. The predominant 3’-5’ exonuclease activity of the T4

DNA polymerase generates 5’-CATTACCATCCC-3’ and 5’-CCAATTCTACCC-3’ overhangs

at either end of the PCR products. Using a similar method but with the addition of dGTP instead,

complementary overhangs were generated on the SmaI restriction enzyme digested pT4P vectors

using T4 DNA polymerase, for directional insertion of the PCR product. The T4 DNA

polymerase-treated PCR product and pT4P vector were then mixed and allowed spontaneous

overhang binding, without an enzyme-mediated ligation step, and then transformed into DH5𝛼

E. coli. Positive clones of the candidate transcripts were then verified by Sanger sequencing.

2.4 Riboprobe synthesis for expression analysis

Primer set AA18 (5’-CATTACCATCCCGCCACCGGTTCCATGG-3’) and PR244F (5’-

GGCCCCAAGGGGTTATGTGG-3’) was used with the templates of cloned transcripts in pT4P

vectors in order to generate PCR products containing one T7 site in the antisense direction to the

3’ ends of the inserted transcripts. Then using a previously published method, antisense

riboprobes were synthesized using T7 RNA-polymerase (Thermo Scientific, Catalog number:

EP0111) for a reaction time of 3 hours (Pearson et al., 2009). During this reaction, 400ng of the

abovementioned PCR product was used as the DNA template. And either digoxigenin (DIG,

Roche, Product No. 11277073910) or fluorescein (Roche, Product No. 11685619910) was used


for RNA riboprobe labeling. A hydrolysis reaction was performed on the synthesized riboprobes

prior to use in whole-mount in situ hybridization (WISH) experiments.

2.5 WISH and dFISH for expression analysis

Whole-mount in situ hybridization (WISH) and double fluorescent in situ hybridization were

performed by adapting previously published methods (Pearson et al., 2009). In both cases,

planarian animals roughly 4mm in length were killed using 5% N-acetyl-cysteine (NAC; Sigma)

in PBS, a mucolytic compound, and then fixed with 4% formaldehyde in PBSTx (formaldehyde:

Sigma; PBSTx: 1× PBS + 0.5% Triton X-100). The fixed animals were treated with a reduction

solution (reduction solution: 50 mM DTT, 1% NP-40, 0.5% SDS, in 1× PBS) and then subjected

to dehydration in 50% methanol solution and 100% methanol solution sequentially (50%

methanol solution: equal volumes of 1× PBSTx and 100% methanol). Both the reduction and

dehydration steps function to permeabilize the fixed animal samples allowing access of

riboprobes, antibodies, and substrates for in situ development. The dehydrated samples were then

treated with a 6% bleach solution (6% H2O2 in Methanol) to bleach pigment present in the

animals to help with in situ pattern imaging. After overnight treatment in 6% bleach solution, the

animal samples were rehydrated using 50% methanol and PBSTx solutions sequentially.

Hybridization of riboprobes was then performed after a 2-hour Prehyb incubation of the animal

samples in riboprobe mix (Hybridization solution: 50% De-ionized Formamide (Roche); 5%

Dextran Sulfate (Sigma); 5× SSC (from 20× stock); 1× Denhardt’s solution (Sigma); 100𝜇g/ml

Heparin (Sigma, from porcine intestine, from 100× stock); 1mg/ml yeast torula RNA (Sigma);

1% Tween-20 (Sigma); 0.6% DTT. Prehyb solution: Hybridization solution without dextran

sulfate, Heparin or DTT. Riboprobe mix: 1:500 to 1:1000 times dilution of DIG-labeled

riboprobe or 1:200 to 1:250 times dilution of fluorescein labeled riboprobes in Hybridization

solution). Hybridization was performed at 56℃ for more than 16 hours and then the animal

samples were washed at 56℃ with each of the following solutions twice, 2× SSC solution, 0.2×

SSC solution and 0.05× solution, for a duration of 30 minutes per wash (2× SSC solution: 2×

SSC + 0.1% Triton-X. 0.2× SSC solution: 0.2× SSC + 0.1% Triton-X. 0.05× SSC solution:

0.05× SSC + 0.1% Triton-X. All using H2O as the solvent). After the SSC washes, the animal

samples were blocked with 5% horse serum plus 5% western blocking reagent (Roche) in MABT

(MABT: 100 mM maleic acid, 150 mM NaCl, 0.1% Tween-20, pH to 7.5 with NaOH). After 2

hours of blocking, the animal samples were then incubated with anti-DIG-AP, anti-DIG-POD or


anti-fluorescein-POD antibodies (All three of the antibodies from Roche. anti-DIG-AP, used at

1:4,000, anti-DIG-POD, used at 1: 500, and anti-fluorescein-POD, used at 1:400 in blocking

solution) overnight at room temperature. 6 times of washes with MABT at 20 minutes each were

performed after the antibody incubation. The animal samples were then ready for signal

detection by colorimetric development or fluorescent development.

For samples with epitope detection by anti-DIG-AP undergoing colorimetric development, a pre-

incubation of 10 minutes in AP buffer was performed (AP buffer: 100 mM Tris, pH 9.5; 100 mM

NaCl; 50 mM MgCl2; 0.1% Tween-20 brought up to volume with 10% polyvinylalcohol solution

(PVA; Sigma P8136)). AP buffer was replaced with Developmental buffer containing the

NBT/BCIP substrate (Development buffer: AP buffer with 4.5𝜇L/mL NBT (Roche) and

4.5𝜇L/mL BCIP (Roche)). The colorimetric development was carried out for a duration of 10

minutes to 3 hours, until an optimal signal to background ratio was achieved. The

colorimetrically developed animals were washed with PBSTx twice and post-fixed with 4%

formaldehyde in PBSTx for 20 minutes. Another two PBSTx washes were performed after the

fixation before the post-fixed samples were treated with 100% Ethanol. Samples were incubated

in 100% Ethanol, which removes the unspecific background staining and turns the NBT/BCIP

precipitate dark blue, for 5-20 minutes until optimal signal to background ratio was achieved.

100% Ethanol was then replaced with another PBSTx wash and the samples were then placed in

80% Glycerol solution overnight, before mounted onto slides for imaging (80% Glycerol

solution: 80% Glycerol; 10mM Tris, pH 7.5; 1mM EDTA). The colorimetric WISH stains

prepared according to the above procedure were imaged on a Leica M165 fluorescent dissecting

microscope. Images were post-processed in Adobe Photoshop. All colorimetric quantification

were performed with the freely available ImageJ software (http://rsb.info.nih.gov/ij/).

Two types of fluorescent developments were performed for samples undergoing fluorescent

development. FastBlue development was performed on samples with epitope detection by anti-

DIG-AP, as previously described (Zhu et al., 2015; Lauter et al., 2011). Samples after MABT

washes were incubated in SB8.2 buffer for 10 minutes (SB8.2 buffer: 100 mM Tris, pH 8.2; 100

mM NaCl; 50 mM MgCl2; 0.1% Tween-20 brought up to volume with 10% polyvinylalcohol

solution (PVA; Sigma P8136)). SB 8.2 buffer was then replaced with FastBlue development

buffer containing the FastBlue and NAMP substrates (FastBlue development buffer: mix equal

volume of 5𝜇L/mL Fast Blue BB (Sigma F3378) and 5𝜇L/mL of naphtol-AS-MX-phosphate


(NAMP, Sigma N5000), for a final concentration of 2.5𝜇L/mL of each substrate). Development

of FastBlue stain was monitored by its blue colorimetric development for a duration ranging

from 10 minutes to 3 hours. The development was stopped as soon as a faint colorimetric pattern

similar to that observed with NBT/BCIP development can be observed under dissecting scope.

In the cases of epitope detection by anti-DIG-POD and anti-fluorescein-POD, the in situ stains

were developed with FITC or Cy3 tyramides. To prepare for the tyramide development, samples

after the MABT washes were first incubated in PBSTx for 30 minutes. Then PBSTx was

replaced with tyramide solution for another 30-minute incubation (Tyramide solution: 1:500

dilution of Cy3 tyramide or 1:2000 dilution of FITC tyramide in PBSTx). After the tyramide

solution incubation, two spikes at 25 minutes apart of H2O2 at a final concentration of 1 part per

10,000 were performed to the samples in tyramide solution. After 45 minutes of the initial spike

of H2O2, the tyramide development was stopped by replacing the tyramide solution with PBSTx.

Another 2 washes of PBSTx for 5 minutes each was performed before the animals were put into

AntiFade mounting media (AntiFade mounting media: 80% Glycerol solution + 1% DABCO


Tyramide development was coupled with FastBlue development or another tyramide

development to produce multicolor fluorescent in situ hybridization stains. Multiple riboprobes

labeled with different epitopes, for instance, DIG and fluorescein, were added to the same

riboprobe mix. The hybridization and first round tyramide development were performed

similarly as in the case of single riboprobe in situ. In the case of a second round development

using FastBlue, anti-DIG-AP is already added during the antibody incubation for the first round

development, thus, the samples after first round tyramide development were incubated in SB 8.2

buffer and proceed as normal for FastBlue development. In the case of a second round

development using another tyramide development, the first round POD-conjugated antibody

activity was quenched by incubating the samples in sodium azide solution for 60 minutes

(Sodium azide solution: 100mM sodium azide in PBSTx). The sodium azide solution was then

washed 5 times with PBSTx for 5 minutes each before the samples were incubated in the next

round of antibody overnight in blocking solution. And the second round tyramide development

proceeded similarly as first round development with a different tyramide. After all developments

are done, samples were placed in AntiFade mounting media. All fluorescently developed

samples, including FastBlue and tyramide stains, were imaged on an Olympus IX81 Quorum


Spinning Disk Confocal with a Hamamatsu C9100-13 EM-CCD camera and a Yokogawa CSU

X1 scanhead. Whole-animal images were stitched together using the application Volocity, and

all images were post-processed in Adobe Photoshop.

2.6 Targeted functional knockdown by RNA interference

RNA interference (RNAi) targeting cloned candidate transcripts was performed as previously

described to knock down their functions in planarian pigmentation and pigment cell lineage

specification (Newmark et al., 2003). RNAi in this study was performed by feeding planarians

with RNAi food in the form of a beef liver paste containing bacteria expressing double-strand

RNAs of the targeted transcript. In order to produce the bacteria culture to make the RNAi food,

pT4P vectors containing target transcript sequence, as described in section 2.3, were transformed

into HT115 E. coli strain, which is capable of RNA synthesis by T7 RNA polymerase when

induced by isopropyl 𝛽-D-1-thiogalactopyranoside (IPTG). In the pT4P vector backbone, there

are two T7 promoters flanking and going in the direction towards the insertion site of the cloned

transcript, which are key to produce complementary strands of the double-stranded RNA

responsible for RNA interference. E. coli bacteria containing the cloned pT4P vectors were

grown to an optical density at 600nm (OD600) of 0.8 and then induced with 1mM IPTG. The

induced bacteria culture was then incubated at 37℃ for another 2 hours, allowing RNA

production by T7 RNA polymerase. Bacteria were then pelleted and mixed with beef liver paste

at a ratio of 333𝜇L of liver per 100 mL of original bacteria culture volume. After thorough

mixing of the beef liver paste and the bacteria pellet, blue food coloring was added to the mixture

at 1 𝜇L coloring /60 𝜇L food. The RNAi food was then aliquoted to 60𝜇L per feeding, which is

sufficient to feed 40 worms in the size range of 2-4mm in length, and stored at -80℃ until use.

The negative control RNAi was prepared in a similar way but with a pT4P vector containing

GFP sequence at the insertion site. During the screening of all genes in this study, RNAi food

was fed to >7 day starved planarians at a frequency of twice per week for 5 weeks, for a total of

10 feedings. Certain phenotypes take fewer feeds and less time to display, and the time points in

figures denote the number of feeds for the specific gene being knocked down as well as days

after the last feeding of RNAi food. For example, 10fd14 denotes 10 RNAi feedings and 14 days

after the last feeding, and 12fd7 corresponds to 12 RNAi feedings and 7 days after the last




3.1 Identified hmbs and alas as planarian pigment cell markers

3.1.1 Selection of candidates and their expression patterns

Analysis of Day 10 depigmented animal and control animal RNAseq data yields 149 genes with

more than 2-fold decreased and 162 genes with more than 2-fold increased expression levels in

the depigmented animals. Of these 311 candidates, 69 down-regulated and 40 up-regulated genes

were cloned based on the descending order of their fold changes. Among the 109 cloned genes,

60 of them give specific WISH patterns. Expression patterns can be categorized as expression in

the gut, in the sub-epidermis with spider morphology (rich in processes extruding from cell

soma), in the sub-epidermis with punctate morphology, in the epidermis with unknown

morphology, in the anterior, in the pharynx, in the para-pharyngeal region, in the neural system

and in the parenchyma with variable expressions across individual animals (Figure 4). Most sub-

epidermal expression patterns (spider and punctate patterns) are generated by genes with down-

regulated expression in depigmented animals. All the variable parenchyma expression patterns

are generated by genes with up-regulated expression in depigmented animals. Other categories of

expression patterns are generated by either down- or up-regulated genes. Up to 3 representative

genes of each category were listed in Figure 3. The expression pattern screen of differentially

expressed genes presented here provides guidance for later functional studies of planarian

depigmentation and repigmentation. In order to establish molecular markers of pigment cells,

genes with distinct sub-epidermal expressions were prioritized for RNAi screening (Figure 4,

rows 2-4). For the identification of regulators of depigmentation and repigmentation, genes with

distinct gut and variable parenchyma expression patterns were prioritized for screening (Figure

4, first and last rows). Many up-regulated genes without distinct wild-type stains were also

screen by RNAi knockdown. The rationale behind screening of these genes is that up-regulated

genes without a distinct expression pattern in normally pigmented animals may still have a

dramatically different expression during, and possibly be required for, depigmentation and

pigment cell regeneration.


Figure 4 Expression patterns of genes with differential expression levels in depigmented animals. Up to 3 representative expression patterns by WISH of each category are listed. The number of genes that fall into each category is displayed in the brackets following category names. Categories of specific expression patterns are gut, sub-epidermal with spider morphology, sub-epidermal with punctate morphology, epidermal (other), anterior, pharynx, para-pharyngeal, neural, ubiquitous, other distinct patterns, and parenchyma variable. For genes in the parenchyma variable category,

two representative images of expression in different individuals are shown. Scale bar: 100𝜇m.


3.1.2 Genes with sub-epidermal expressions and pigment cell-like morphologies

The expression patterns of spider and punctate categories of genes resemble the anatomical

location of pigment cells as previously proposed and are further characterized to determine

whether they are marking pigment cells (Palladini et al., 1979a). Spider gene category includes

an hmbs homolog (spider-1). Hydroxymethylbilane synthase (HMBS), encoded by hmbs, is an

enzyme responsible for the third step of heme synthesis pathway catalyzing the production of

hydroxymethylbilane from porphobilinogen (Grandchamp et al., 1987). Another spider gene is

an alas homolog (spider-2). Aminolevulinic acid synthase (ALAS), encoded by alas, is the

enzyme responsible for the first step of the same heme synthesis pathway as HMBS, responsible

for the conversion of succinyl-CoA to d-Aminolevulinic acid (Bishop, 1990). HMBS functions

in the cytoplasm while ALAS is usually found in the mitochondria (Grandchamp et al., 1987;

Bishop, 1990). Punctate genes category includes an ans homolog (punctate-1). L-Asparaginase,

encoded by ans, catalyzes the deamination reaction from L-asparagine to L-aspartate (Cantor et

al., 2009). A homolog of sds (punctate-2) also belongs to the punctate category. L-serine

dehydratase/L-threonine dehydratase, encoded by sds, is responsible for the catalytic activity of

converting L-serine and L-threonine to pyruvate and 2-oxobutanoate, respectively (Ogawa et al.,

1989). Both the punctate markers and spider markers display distinct expression patterns

throughout the entire animal in the sub-epidermal layer (Figure 5A). The expression of both

types of markers is quite uniform within the dorsal or ventral surface. However, the dorsal

surface has a higher density of both markers over the ventral surface. spider-1, hmbs, is

expressed at a density of 161±44 cells/mm2 in the dorsal surface (n = 6). Whereas punctate-1,

ans, is expressed at a density of 1137±71 cells/mm2 in the dorsal surface (n = 6). One exception

to the generally uniform expression of spider and punctate markers is the lack of expression in

the eye region. No hmbs expression can be detected in the patches of cells opposing the planarian

eye cups, which include photo-sensory neurons (Figure 5B). This is a logical expression pattern

for potential pigment cell markers, because planarian lacks pigmentation in the photo-sensory

region, allowing unhindered light passage. It is also observed that cells expressing spider

markers and punctate markers are not located in the outermost layer of the planarian epidermis

(Figure 5B, C). Individual cells labeled by punctate marker ans and sds display small and round

morphology. Individual cells labeled by hmbs and alas are observed to have cytoplasmic

processes extending from the cell soma. All the morphological observations about cells labeled


by spider cell markers agree with previously published data on planarian pigment cells, thus are

suitable candidates for pigment cell markers (Figure 5D) (Palladini et al., 1979a, 1979b).

Figure 5 Expression patterns of spider and punctate genes resemble pigment cell morphology. A) Spider and punctate genes are expressed throughout the surface of planarian animals. The dorsal planes of animals are shown. B) Details of hmbs expression in the anterior region. Individual hmbs positive cells have extending processes from the cell soma. A gap exists between hmbs expression and the edge of the dorsal plane, suggesting expression of hmbs in the sub-epidermal layer. Patches opposing eye pigment cups lack hmbs expression. C) Details of ans expression in the posterior region. Individual ans positive cells have small and round morphology. D) Illustration of pigment cell morphology in axial cross section. E, epidermis. BM, basal membrane. MF, muscle fibre. P, parenchyma. Black dots in the pigment cell indicate pigment granules. All expression patterns stained by WISH. Scale bar: 100𝜇m.

3.1.3 Expression of spider markers correlates to pigment status of planarian tissue

After morphologically establishing spider and punctate markers as suitable candidates for

pigment cell markers, I next tested whether their expression patterns were affected during

depigmentation, as predicted if they are bona fide pigment cell markers. Planarians roughly 3mm

in length were exposed to visible light in the LED light exposure set up for 7 days. The animals


Figure 6 Expression of spider and punctate genes directly correlates to the pigment status of planarian tissue. A) Live image of a light-exposed planarian animal. (>8 days, completely depigmented) B) Live image of a regenerating planarian. 7 days after amputation. Newly regenerated head and tail (blastema) are mostly not pigmented. C) Normal expression of hmbs (spider-1), pepck (spider-3) and ans (punctate-1) in WT (control) diminishes in 7-day light exposed animals (mostly depigmented). D) hmbs is lowly expressed in newly regenerated blastema at 5-day post amputation. F) Details of limited expression of hmbs in 5-day blastema. All expression patterns stained by WISH. Panels D and F are courtesy of Stephanie

Hallows. Scale bar: 100𝜇m.

were almost completely depigmented at the end of this treatment (Figure 6A). I then performed

WISH on these light exposed animals and determined the expression level of spider and punctate


markers. The expression of punctate markers is undetectable while spider marker expression is

greatly reduced (n ≥ 3 for each WISH stains of both conditions) (Figure 6C). The positive

correlation between the pigment status of planarians and the expression of spider and punctate

markers supports these genes as pigment cell markers.

Another method to generate unpigmented planarian tissue is by amputation (Figure 6B). The

newly regenerated blastema after amputation is unpigmented and it takes about 2 weeks before

the pigment status is restored to a similar level to that of the old body part. With assistance from

the Pearson Laboratory technician, Stephanie Hallows, we performed WISH of spider markers in

newly regenerated blastema at 5 days post amputation. Only a handful of cells displayed

expression of the spider-1 marker in both the anterior and posterior blastema (n > 10, counting

head, trunk, and tail fragments) (Figure 6D, F), further supporting the positive correlation

between the expression of spider markers and pigment status of the planarian tissue.

3.1.4 hmbs is functionally required for pigment production and maintenance

For a marker with high specificity, its expression should be exclusive within the pigment cell

population only. And logically, I hypothesized that the exclusive expression of these genes in

pigment cells are crucial for pigment cell functions, for instance, pigment production and

maintenance. Thus, I performed functional knockdown of several punctate and spider genes by

RNAi and observed changes in planarian pigmentation. After 12 feeds of RNAi, functional

knockdown of punctate genes did not show any effect on planarian pigmentation or the recovery

of pigmentation after light exposure (n = 28). On the other hand, when spider-1, hmbs, was

functionally knocked down by RNAi, the pigmentation of hmbs RNAi animals became much

lighter than that of control animals (n = 35) (Figure 7A). spider-1 RNAi animals did not display

any apparent health or behavioral defects. Although the treated animals are much lighter in

pigmentation, residual pigmentation with similar expression morphology to that of control

animals can still be observed.

I next tested the functional requirement of spider-1 during the repigmentation of blastema after

amputation. Control and spider-1 RNAi worms after 12 feeds were amputated at 7 days after the

last feeding of RNAi food and allowed to regenerate for 30 days. Roughly 7 days is required for

planarians to regenerate new head and tail tissue after amputation and morphallax to similar body

proportions as before amputation (Newmark and Sánchez Alvarado, 2002). In this experiment,


Figure 7 Functional requirement of hmbs in pigmentation maintenance and repigmentation. A) Representative live images of animals after 12 feeds of RNAi food and 7 days after the last feeding. Control animals display normal pigmentation while hmbs RNAi worms are no longer pigmented. B) Representative live images of animals after similar treatment as in A), but imaged with amputation performed at 7 days post RNAi feedings and after 30 days of regeneration. In control animals, pigmentation and boundaries of clearing around eyes are established in newly regenerated tissue. In hmbs RNAi animals, pigmentation is not established in neither the old body parts nor the newly regenerate tissue.

an extensive 30-day regeneration period allowed the amputated animals to fully regenerate their

pigment cells in the new tissue and produce body pigment. In the control animals, I observed that

the newly regenerated head and tail tissue have recovered substantial pigmentation, although still

slightly lighter than the old body parts, at the end of the regeneration period (n = 4, counting

trunk fragments only) (Figure 7B). I also observed that the boundaries of the characteristic

pigment-less patches in the eye region have been re-established, due to sufficient body pigment

deposition in the surrounding tissues. In the spider-1 RNAi worms, however, the newly

regenerated head and tail contained no detectable recovery of body pigment (n = 6, counting

trunk fragments only) (Figure 7B). There is no discernible boundary of pigmentation between the

eye region and the surrounding tissues to be detected, either. Furthermore, even the old body

parts remained largely unpigmented. These results suggest that hmbs is responsible for the

maintenance of planarian body pigmentation during homeostasis and for the pigment production

in newly regenerated tissues, supporting hmbs as the planarian pigment cell marker functionally.

3.1.5 hmbs is co-expressed with other spider markers and kmo

After establishing hmbs as planarian pigment cell markers with morphological and functional

evidence, I then determined its co-expression with several other spider and punctate markers

identified in this study. By performing dFISH, I observed that less than 7% of ans positive cells


also express hmbs (Figure 8A). This result indicates that the two genes are mostly not co-

expressed. On the other hand, dFISH of hmbs and alas shows that 97% of alas expressing cells

also express hmbs (Figure 8B). Not only does the relative position of the dFISH signals of the

two genes overlap, but also do the overlapping dFISH signals have similar morphology,

indicating the two genes are co-expressed.

I then performed dFISH to determine if punctate markers are co-expressed with each other. Due

to the low yield of fluorescent signals in Tyramide Signal Amplification (TSA), however, no

usable data was obtained. Several reasons can account for the low detectability of punctate

markers by TSA. The punctate gene expression levels are relatively low and the accessibility to

them by RNA probes might be hindered, depending on the subcellular localization. In addition,

POD-conjugated antibodies are inherently ineffective at amplifying fluorescent signals from

weak RNA probes. The enzymatic activity of POD-conjugated antibodies usually diminishes

after 45 minutes of spiking by H2O2, making detection of punctate gene expression using POD

extremely difficult. I then tested the method of using two rounds of alkaline phosphatase

reactions with different fluorescent substrates, to sequentially detect the two punctate genes. This

attempt also failed, due to the intense background generated by HNPP substrates used in this

approach. I was able to, however, perform ans in situ alone (n = 5), sds in situ alone (n = 5), and

in situ of ans and sds combined (n = 5) (with DIG-labeled riboprobes of both transcripts added to

the same riboprobe mix) in three separate samples (Figure 8D). By counting cells labeled by the

above riboprobe mixes, I discovered that ans (2472 ± 530 cells/mm2) and sds (2764 ± 128

cells/mm2) positive cells are of similar density to each other (Figure 8D). Furthermore, when two

probes are used together, there is no increase in the density of labeled cells comparing to that

detected by using single probes alone (2484 ± 293 cells/mm2) (Figure 8E). In addition, the

WISH development of double probes were much quicker and stronger within a similar timeframe

than that of single probes (Figure 8D, top left panel). These results suggest that the two punctate

markers, ans and sds, are co-expressed in the same population of cells. Note that the density of

punctate cells calculated here is the result of counting WISH stains on the epi-fluorescent

microscope of both the dorsal and ventral sides of the samples, thus, the density is roughly

doubled of that calculated from confocal images of the dorsal surface only.

The transcript of a gene in the tryptophan metabolism pathway, kynurenine 3-monooxygenase

(kmo), was cloned by my collaborator, Dr. Jason Pellettieri (Croucher et al., 2013; Linzen,


Figure 8 Co-expression analysis of spider and punctate markers. A) Most hmbs or ans positive cells do not co-express with each other. B) hmbs and alas are highly co-expressed. C) hmbs and kmo are highly co-expressed. White arrows indicate a lack of co-expression in A), and cells that are co-expressing markers in B) and C). Approximate locations on the dorsal plane of the animals imaged were shown to the right. D) ans and sds expression are similar in cellular population and morphology. WISH with combined ans and sds probes generates similar expression pattern to individual probes without apparent changes in population or morphology. E) The density of positively stained cells in each

condition shown in D). Error bar: Standard deviation. Scale bar: 100𝜇m.


1974). kmo shows an expression pattern similar in morphology and density to that of other spider

markers (Figure 8C). I then performed dFISH between kmo and hmbs and observed that hmbs is

co-expressed in 96 ± 4.2% of kmo positive cells (Figure 8C). The tryptophan metabolism

pathway, in which kmo functions, is key to the production of ommochromes, one of the major

types of pigments found in the animal kingdom. Reports of studies in mosquitos and silk worms

show that kmo is required for proper pigmentation of the eyes and silk color, in either organism,

respectively (Han et al., 2003; Osanai-Futahashi et al., 2012). I also tested the expression pattern

of 6 transcripts involved in tyrosine metabolism, a pathway involved in melanin production, and

did not find any spider cell-like expression patterns. In summary, the co-expression of kmo and

hmbs in pigment cells strongly suggests that ommochrome is one of the responsible pigments for

the body pigmentation of S. mediterranea. This result agrees with previous studies on purified

planarian body pigment using spectrophotometry in the UV region (Hase et al., 2006).

3.1.6 Pigment cells show inward migration toward gut during depigmentation in response to light exposure

Using the established pigment cell marker hmbs, a basal timeline for planarian pigment cell loss

during acute light exposure was established. I found that planarian pigment cells are lost

gradually over the course of 10 days under white LED light exposure. Animals were mostly

depigmented after 7 days of light exposure (Figure 9A). A 12-day light exposure schedule was

used in this experiment to ensure a complete depigmentation. The population of planarian

pigment cells, marked by hmbs, gradually reduces over the time course of the light exposure (n

≥ 5 for each time point) (Figure 9B). However, the greatest change in pigment cell population

was observed after Day 5 during the light exposure. The disappearance of hmbs positive cells is

slightly slower than the disappearance of pigmentation. The most dramatic change in the body

pigmentation status is observed before Day 5 (Figure 9A). This discrepancy between

pigmentation and pigment cells disappearance is likely due to the hypothesized mechanism that

pigments within individual pigment cells are bleached by light exposure before generating ROS

and leading to cell death. At the end of the light exposure time course, hmbs expression is not

detectable in the majority of planarian tissues (Figure 9B, right panel). Throughout the entire

depigmentation timeline, it is also observed that the head and tail regions are the first to become

depigmented while the para-pharyngeal region remains pigmented for longer durations (Figure


Figure 9 Pigment cells migrate towards gut during depigmentation. A) Representative live images of planarians during light exposure induced depigmentation time course. B) Pigment cells, marked by hmbs, decrease during the time course. Axial cross sections show no hmbs expression in the parenchyma in Day 0 (un-exposed) animals. But hmbs expression is increasingly detected in the parenchyma near the gut as the time course progresses. In the meantime, expression in the sub-epidermal layer decreases. C) Axial cross sections of planarians show the appearance of pigment/pigment cells in the gut in Day 5 light exposed animals.


9B). The difference of depigmentation across different regions of the body is evidenced both by

the direct observation of body pigmentation in live worms and by labeling pigment cells with

hmbs riboprobes.

During the light exposure time course, I have observed darkening of the gut in the treated

animals, especially at 5 days after the start of the light exposure. Comparing the axial cross

sections of depigmenting planarians with those of the animals kept in dark, there is an evident

increase of the accumulation of pigment in the gut of depigmenting planarians (n = 5, the number

of animals used to generate cross sections) (Figure 9C). In certain cross sections of the

depigmented animals, concentrated spots of pigmentation can be observed in addition to smears

of pigment in the gut region, suggesting possible excretion of pigment cells instead of pigment

alone (Figure 9C, right panel). In order to determine if pigment cells are indeed removed through

gut excretion during light exposure, I performed experiments to detect the changes of hmbs

positive cells in axial cross sections (n ≥ 4, the number of animals used to generate cross

sections for each time point). In control animals without light exposure, hmbs positive cells are

only observed in the sub-epidermal region (Figure 9B, left panel). At Day 5 of light exposure,

however, a substantial amount of hmbs positive cells can be observed ectopically in the central

parenchyma of the planarians, a region in proximity to the gut (Figure 9B, centre left). On day 7

of light exposure, an overall reduction in the number of hmbs positive cells can be observed

(Figure 9B, centre right). And among the remaining hmbs positive cells, the majority are

positioned ectopically in the central parenchyma region close to the gut. The ectopic expression

of hmbs expression is exacerbated in the Day 10 animals, where in many cross sections the

central parenchyma region becomes the only location of hmbs expression (Figure 9B, right). In

summary, the reduction of pigment cell population, marked by hmbs, and the observation of

inward migration of pigment cells suggest that an active removal of dying pigment cells through

gut excretion likely happens during light exposure. These findings agree with previous apoptosis

analysis using TUNEL by my collaborator, Dr. Jason Pellettieri (Figure 2B, C).

3.1.7 Pigment cells display fast regeneration timeline during repigmentation

After 12 days of light exposure, the depigmented animals were placed in dark allowing for

repigmentation. Whole worm samples were collected during repigmentation (n ≥ 5 for Day 1.5 –

8.5, n = 2 for Day 12.5). And the regeneration of pigment cells was analyzed using hmbs marker.


The time point at the conclusion of light exposure was counted as time point zero for the

regeneration time course. Sporadic expressions of hmbs can be observed in animal samples

collected at as early as Day 1.5, suggesting rapid pigment cell regeneration (Figure 10B). A

gradual increase of pigment cell population can be observed from this time point onwards. As

early as 7 days into the regeneration, pigmentation spots can be observed on live animals with

expected pigment cell morphology. After more than 12 days of repigmentation, pigment cells are

regenerated throughout the entire body of the previously depigmented animals (Figure 10B,

right). Despite this rapid regeneration of pigment cells, the pigmentation status of planarians at

this time point is still very low and sporadic (Figure 10A). Only patches of pigmentation can be

observed while most parts of the body remained unpigmented. This difference in the rate of

pigment cell regeneration and repigmentation suggests the possible lengthy process of pigment

production taken by the newly regenerated pigment cells. During the two weeks of

repigmentation analyzed in this study, pigment cells did not display any preferences or any

differences of the regeneration rates in the head, trunk or tail regions (Figure 10A, B).

Figure 10 Rapid regeneration of pigment cells during repigmentation. A) Representative live images of planarians during repigmentation time course following depigmentation. B) Pigment cells, marked by hmbs, are rapidly regenerated during repigmentation. Regeneration of pigment cells precedes the appearance of pigmentation.


3.1.8 Decreasing levels of co-expression between punctate and spider markers as repigmentation progresses

In control animals, punctate cells labeled by ans have a density of 1137 ± 71 cells/mm2 in the

dorsal surface, much higher than the density of 164 ± 44 cells/mm2 for pigment cells labeled by

hmbs (Figure 11A, B). Only 6.99% (n = 6) of all ans positive cells in control animals also

express hmbs (Figure 11A, B). Considering the resolution of the imaging equipment and

technique used, false-positive likely accounts for a substantial portion of the co-expression

observed. The convincing evidence of substantial co-expression between ans and hmbs comes

from dFISH experiments at early time points during repigmentation. At Day 1 and Day 3

repigmentation, 78.2% (n = 3) and 64.6% (n = 4) of ans positive cells co-express hmbs,

respectively (Figure 11A, B). In two samples at Day 1 repigmentation, almost all hmbs positive

cells are co-expressing ans (32 of 33 hmbs+ cells for Individual 1, and 13 of 13 for Individual 2).

This observation strongly suggests ans expression as a key feature of newly differentiated

pigment cells. This high level of co-expression gradually drops to a moderate level of 27.4% (n =

2) and 29.7% (n =1) at Day 8 and Day 11 during repigmentation, respectively (Figure 11A, B).

The repigmentation time course also shows that across all time points and all individual animals

tested, ans always has a wider expression than hmbs (Figure 11A, B). In summary, the

percentage of co-expression is highest at the earliest time point of repigmentation following

punctate and spider cell depletion. And the co-expression decreases as punctate cells are more

rapidly regenerated than pigment cells.


Figure 11 Co-expression of spider and punctate genes decreases along repigmentation time course. A) In control animals, most ans positive cells do not co-express hmbs. High level of co-expression exists early during repigmentation. At later time points, the percentage of co-expression drops. B) Magenta: density of ans positive cells in the dorsal plane. Green: density of hmbs positive cells in the dorsal plane. Gray: Percentage of ans positive cells that are co-expressing hmbs. Error bar: Standard deviation. Scale bar: 100𝜇m.


3.2 Identified regulators of depigmentation and pigment cell regeneration

3.2.1 hmbs is not required for pigment cell survival

Earlier, I showed that RNAi knockdown of hmbs leads to depigmentation of planarians and

argued the finding to be a functional evidence to establish hmbs as a molecular marker for

pigment cells. I further analyzed hmbs RNAi knockdown to study its effect on pigment cells and

related cells (n ≥ 8 for each of the stains). It is observed that after 12 feedings of hmbs RNAi, the

expression level of hmbs transcripts is no longer detectable by WISH (Figure 12A). On the

contrary, the expression level and the number of cells expressing alas is similar to that in the

control animals (Figure 12A). As mentioned earlier, alas is a redundant marker to hmbs that also

labels pigment cells (Figure 8B). I also tested the effect of hmbs RNAi on the punctate marker

ans, which potentially marks pigment cell progenitors, but found that the expression level and

the population of positive cells are not affected (Figure 12A). This experiment demonstrates that

although hmbs is required for pigment production and maintenance, it is not required for the

survival of pigment cells or their potential progenitor cells.

3.2.2 Presence of pigment is required for light-induced pigment cell loss

Several previous studies suggested that various types of animal pigment can produce reactive

oxygen species (ROS) in response to light exposure. At high levels, ROS is detrimental to

cellular functions and can lead to cell death (Shen et al., 2015; Pospíšil et al., 2014). I

hypothesized that ROS produced by planarian pigment causes pigment cell death and removal

during light exposure. It has been established that hmbs RNAi leads to pigment loss without loss

of pigment cells. Thus, working with hmbs RNAi animals, I investigated the effects of light

exposure on pigment cells without the presence of pigment. Animals were fed with control and

hmbs RNAi food for 10 times. At the conclusion of RNAi feedings, the hmbs RNAi animals

appeared almost unpigmented while the control animals displayed normal body pigmentation (n

= 10 for both). The RNAi animals were then treated with light exposure for 9 days, a period that

is sufficient to completely depigment the control animals. The hmbs RNAi animals remained

unpigmented throughout the light exposure treatment. I then determined the population of

pigment cells in the light exposed animals using the secondary pigment cell marker alas. As

predicted, almost no pigment cells were detectable by WISH in the control RNAi animals


Figure 12 hmbs is not required for pigment cell survival and its RNAi offers protection from light exposure. A) WISH analysis of pigment cell lineage markers in control and hmbs RNAi animals. hmbs RNAi results in undetectable levels of hmbs transcripts by WISH, but does not affect the survival of pigment cells, marked by alas, or pigment cell progenitors, marked by ans. B) hmbs RNAi animals retain a population of pigment cells, marked by alas, after 9 days of depigmentation,

while control animals lose all pigment cells. Scale bar: 100𝜇m.

(Figure 12B). In the hmbs RNAi worms, despite appeared as unpigmented, the expression level

of alas remained high in individual cells and the population of alas positive cells remained at a

comparable level to untreated animals (Figure 12B). These results support the hypothesis of ROS

generated by body pigment in response to light being the cause of planarian depigmentation and

pigment cell loss after light exposure.


3.2.3 epimerase positively regulates depigmentation in response to light

Transcript SmedASXL_011154, encoded by a gene that is homologous to epimerase, is

exclusively expressed in the triclad-gut of planarians (Figure 4, top row, left panel). Epimerases

are isomerase enzymes responsible for the inversion of the stereochemistry of biological

molecules. They are reported to be involved in a wide variety of biological processes, including

sugar metabolism and protein sialylation, which are important for signal transduction and cell

surface adhesions (Daude et al., 1995; Keppler et al., 1999). This gene is observed to be 5 times

down-regulated in depigmented animals according to the RNAseq data. After 10 feeds of RNAi,

I performed light exposure treatment on the RNAi animals. At Day 5 of the light exposure

treatment, the majority of the pigmentation of the control animals has disappeared (Figure 13A).

Although a limited population of pigment cells generally remains at this time point (Figure 9B),

the worms appeared mostly colorless (Figure 13A). On the other hand, the epimerase RNAi

worms at Day 5 depigmentation retained most of their body pigmentation (Figure 13A). The

difference of depigmentation between epimerase and control RNAi worms are obvious enough

to be noticed by observing whole plates of animals with the naked eye. Under the dissecting

scope, epimerase RNAi animals retained pigmentation in the majority of their bodies, with

limited depigmentation in the head and tail region only (Figure 13A). The results from this

experiment suggest that epimerase is required for planarian depigmentation in response to light


3.2.4 draper positively regulates depigmentation in response to light

Transcript SmedASXL_006481, encoded by a gene that is homologous to draper in Drosophila

Melanogaster, or ced-1 in Caenorhabditis elegans, has a weak, non-specific and ubiquitous

expression throughout the entire planarian body. ced-1 and draper are reported to be responsible

for the removal of cell corpses by engulfment (Zhou et al., 2001; Freeman et al., 2003). In C.

elegans, ced-1 encodes a transmembrane receptor which recognizes the signal presented by

CED-7 ABC transporter on the surface of dying cells (Zhou et al., 2001). ced-1 function is

required for the engulfment of dying cells, and the process of cell corpse removal itself promotes

cell death (Zhou et al., 2001). draper, the Drosophila homolog of ced-1, is exclusively expressed

in glial cells and macrophages (Freeman et al., 2003). During Drosophila neural development,

draper is responsible for the engulfment and clearance of apoptotic neurons by glial cells

(Freeman et al., 2003). According to RNAseq data, the planarian homolog of draper only shows


Figure 13 RNAi phenotypes of epimerase, draper and fst. A) epimerase RNAi leads to decreased rate of depigmentation. B) draper RNAi leads to decreased pigment cell removal during light exposure. C) draper RNAi leads to the formation of pigment-less patches on planarian skin. D) fst RNAi leads to increased rate of pigment cell regeneration during repigmentation. E) fst RNAi leads to increased rate of pigment cell production during homeostasis. Percentages of increase in the head, trunk and tail regions are shown in the right panel. Scale bar: 100𝜇m.


a moderate 1.3-fold downregulation in the depigmented animals. I tested its effects on light-

induced depigmentation to determine whether it is required for the clearance of dying pigment

cells. I performed 10 feeds of RNAi and then placed the animals under light exposure treatment.

After 6 days of light exposure treatment, the control animals had no pigment cells remaining as

detected by marker hmbs (n = 5) (Figure 13B). At the same time point, however, draper RNAi

worms still had large numbers of pigment cells in the sub-epidermal layer (n = 6) (Figure 13B).

The impaired pigment cell clearance during light exposure in draper RNAi worms is not biased

toward the head, trunk or tail regions. Locally, however, the distribution of remaining pigment

cells is inconsistent, with randomly distributed pigment cell-free patches and pigment cell-

concentrated areas.

3.2.5 Disruption of draper function leads to formation of wrinkles on epidermis and ectopic expressions of lineage markers

In addition to the defective pigment cell removal caused by draper RNAi, the treated animals

also display wrinkles on their skin tissues. Specifically, during the post-feeding homeostasis,

wrinkles were formed across the entire epidermis, with uniform severity from head to tail. In

addition, draper RNAi worms display pigment-less patches at random locations of the animal

body. A closer look at these patches reveals that they are uprising bumps on the skin. I confirmed

the absence of pigment cells in the pigment-less patches by performing WISH of hmbs on 14

days post-feeding homeostatic RNAi worms (n = 5). In comparison to control, the tissue bumps

in draper RNAi animals were completely devoid of hmbs positive cells (Figure 13C). The

cellular composition in these tissue bumps is unclear and their relation to the defective pigment

cell removal during light exposure is not understood. To address these questions, I tested the

expression of epidermal lineage markers in 10-feed RNAi animals after 14 days of homeostasis

following the last RNAi feed (n ≥ 4 for intact animals of each condition; n = 2, the number of

animals used for both types of axial cross sections). In control animals, piwi-1, a stem cell

marker, is solely expressed in the parenchyma and is absent from the epidermis marked by vim-1

(Figure 14A). The stem cell compartment labeled by piwi-1 is clearly separated from the

epidermal cells, with a gap constituted of other cell types, as shown in axial cross sections of the

dFISH samples (Figure 14A). Early progeny cells in the epidermal lineage, marked by prog-2,

are situated closer to the epidermis than piwi-1 cells, but are absent in the outermost layer of the

planarian tissue, as evidenced by the lack of prog-2 expression at the edges of dorsal planes


(Figure 14B). In the draper RNAi animals, however, the compartmentalized expression of both

piwi-1 and prog-2 is disrupted. piwi-1 and prog-2 positive cells can be detected in close

proximity to vim-1 cells and at the outermost edges of dorsal planes (Figure 14A, B). In axial

cross sections, piwi-1 positive cells show smaller gap with vim-1 cells, and ectopic expressions

of piwi-1 can be detected directly in the epidermis (Figure 14A). It is also observed that the

consistency of epidermis is disrupted in draper RNAi animals. The epidermis is filled by ridges

and ruptures with ectopic expressions of other lineage markers instead of the distinct smooth and

continuous epidermis marked by vim-1 in control animals. These results suggest that draper is

required for the maintenance of the compartmentalization of gene expressions in planarians. The

loss of planarian tissue integrity is likely contributing to the impaired pigment cell removal

during light exposure in draper RNAi worms. However, it is unclear if this function of draper is

related to its cell corpse removal function reported in other organisms (Zhou et al., 2001;

Freeman et al., 2003).

3.2.6 follistatin negatively regulates pigment cell regeneration and homeostasis

Transcript SmedASXL_012581, encoded by a gene that is homologous to follistatin (fst), has

sparse expression throughout the intact planarian and at the anterior pole (Gaviño et al., 2013;

Roberts-Galbraith et al., 2013). Follistatin is a bioneutralizer of ligands in the TGF-𝛽

superfamily, including activins (Knight, 1996; Roberts-Galbraith et al., 2013). TGF-𝛽 and BMP

ligands bind to cell surface receptors which then directs SMAD proteins to regulate transcription

of downstream effector genes (Rahman et al., 2015; Itoh et al., 2014). fst, as a modulator of

TGF-𝛽 signaling pathway, is important for an array of biological processes, including stem cell

pluripotency maintenance and differentiation (Itoh et al., 2014). Previous studies in planarians

demonstrated that fst displays robust expression near wound sites and its function is required for

regeneration following amputation (Gaviño et al., 2013). RNAi knockdown of fst results in

failure of blastema formation after amputation (Gaviño et al., 2013). Here, I performed

experiments to determine the functional requirement of fst in the regeneration of pigment cells

following light-induced depigmentation. Animals were fed with control or fst RNAi food for 10

times and treated with 10 days of light exposure. After the depigmentation, the worms were

placed in dark allowing repigmentation. Whole worm samples were collected for detection of

pigment cells by hmbs riboprobe. The control animals displayed a normal extent of pigment cell


Figure 14 Ectopic expression of lineage markers in draper RNAi condition. A) Compartmentalized expression of stem cell marker, piwi-1, is disrupted in draper RNAi animals. Arrows indicate ectopic piwi-1 expressions and ruptures of the epidermis, marked by vim-1. B) Early progeny cells, marked by prog-2, are ectopically

expressed in the epidermis in draper RNAi animals. Arrows indicate ectopic expressions. Scale bar: 100𝜇m.


regeneration at Day 8 of repigmentation (n = 6) (Figure 13D). The fst RNAi worms are evidently

more pigmented by viewing the whole plate of animals with the naked eye. Using WISH, a much

larger population of regenerating pigment cells were detected in these animals (n = 4) (Figure

13D). This proliferative phenotype is in contrast to the previously published results

demonstrating the failure of regeneration under fst RNAi conditions. To test whether fst RNAi

affects pigment cell proliferation during homeostasis as well, I collected whole worm samples

after 14 days of homeostasis following the same RNAi treatment. Comparing to the control

animals, the fst RNAi worms displayed a significant increase of pigment cell density across the

entire body, ranging from 14.9% increase in the trunk region to the 54.1% increase in the neck

region (n = 3 for both control and fst RNAi) (Figure 13E). These results suggest that follistatin

negatively regulates pigment cell regeneration and homeostasis.



My initial molecular characterization of pigment cell lineage in planarians presented a novel

model to study regeneration and ASC regulation. The light-induced depigmentation allows

investigation in the regeneration of specific tissue types without introducing physical injuries. In

addition, the kinetics of pigment cell and progenitor cell regeneration demonstrate distinct modes

of regulations between different stages of lineage specification. Finally, investigations of gut

excretion mediated pigment cell removal identified regulators of potential cell corpse engulfment

mechanism during the depigmentation process. Taken together, these findings provide new

avenues toward future studies in ASC regulation and gut excretion in planarians.

4.1 Identified markers, and established pigment cells as a novel model for ASC and regeneration study

As hypothesized, I have established several genes with reduced expressions in the depigmented

animals as molecular markers for planarian pigment cells. The expression patterns of pigment

cell markers, hmbs and alas, show a high resemblance of previously published pigment cell

morphology (Palladini et al., 1979a). A direct correlation between the expression pattern and the

pigment status of planarian tissue also supports these genes as pigment cell markers. Finally, the

functional requirement of hmbs in the maintenance and production of planarian body pigment

further supports it as a pigment cell marker. The establishment of these molecular markers made

possible to further study the lineage specification and regeneration of pigment cells in planarians.

Combining with the method of pigment cell ablation by acute light exposure, this new model

allows one to dissect the specific regulators of lineage commitment and to investigate the

mechanisms regulating regeneration, without the introduction of physical injuries.

Important insights of the planarian body pigment composition were gained by co-expression

analysis of pigment cell marker hmbs with other related genes. The lack of epidermal expression

patterns of tyrosine metabolism pathway genes suggests that it is unlikely for melanin to be the

planarian body pigment. On the other hand, hmbs highly co-expresses with a key enzyme

encoded by a kmo homolog in the tryptophan metabolism pathway. kmo is responsible for the

synthesis of ommochrome, the pigment that is responsible for the color of silk worm cocoons

and mosquito eyes (Linzen, 1974; Osanai-Futahashi et al., 2012; Han et al., 2003). In addition,

my preliminary results show that kmo RNAi indeed leads to decreased pigmentation, although


not as profound as hmbs RNAi. This observation provides a direct link between ommochrome

synthesis and planarian body pigmentation. Considering this link with ommochrome, however, it

is unclear why the function of HMBS is highly required for planarian pigment maintenance.

Both hmbs and alas encode key enzymes in the heme synthesis pathway. But heme is not a direct

component of ommochrome synthesis. Heme is, however, a cofactor required for the enzymatic

activity of TDO and IDO, the enzymes responsible for the initial step of ommochrome synthesis

(Meng et al., 2014; Forouhar et al., 2007; Huang et al., 2013). It is unclear why the perturbation

of heme synthesis has a greater impact on pigmentation than the perturbation of ommochrome

synthesis itself. It is possible that planarian body pigment consists of ommochromes in addition

to some other types of pigments, including porphyrins, in which heme is a direct component.

Further characterization of the composition of body pigment could be key to the understanding

of planarian pigment cells. Additional biochemical assays are necessary to address these

questions, including testing for the presence of cytochrome P450 and cytochrome c in planarian

pigment cells.

Pigment cells marked by hmbs were also observed to migrate inwards toward close proximity of

the gut during light exposure. This observation agrees with the previous findings by my

collaborator that planarian gut darkens and shows increased level of apoptosis during

depigmentation. The unique gut excretion phenomenon awaits further investigation to determine

their possible roles as an active mechanism utilized by planarians for removal of detrimental

cells and contributing to their cancer-free immortal lifestyle. During the repigmentation time

course, I discovered a swift regeneration response of planarian pigment cells, marked by hmbs.

Interestingly, it is known that pigment cells have a low turnover rate. One evidence comes from

the retaining of proper pigmentation in lethally irradiated worms, who possess no stem cells to

produce new pigment cells. The swift regeneration response suggests that a possible feedback

mechanism exists between pigment cells and ASCs allowing the increased level of pigment cell

differentiation in response to reduction of its population size. The punctate markers ans and sds

show a high level of co-expression in pigment cells during early repigmentation. The punctate

cells display a much faster regeneration kinetics, and a population expansion outside of pigment

cells is observed in late repigmentation. These observations suggest that punctate cells are likely

progenitors to mature pigment cells. A few possible explanations can accommodate the different

observations of spider and punctate cell regenerations (Figure 15). First, ans positive cells pause


Figure 15 Model for pigment cell lineage proliferation. A) During homeostasis, ASCs (marked by piwi-1) constantly differentiate into pigment progenitor cells (ans) leading to progenitor accumulation. ans positive cells contribute to pigment cells (marked by hmbs, alas and kmo) and other related cell types in a balanced fashion. B) During early repigmentation, a depletion in pigment cell population signals pigment cell progenitors to rapidly differentiate into mature pigment cells. In the meantime, the progenitor cell pool is depleted due to the rate limiting differentiation step from ASCs to progenitors. In addition, differentiation from progenitors to pigment cells is prioritized over other related cell types.


at the progenitor stage and accumulate due to the slow turnover rate of pigment cells (Figure

15A). After depigmentation, ans positive cells are swiftly turned into mature pigment cells as an

immediate response of regeneration (Figure 15B). However, the step of differentiation from

ASCs to progenitors is likely rate-limiting, thus leading to a depletion of progenitor cell pool

during the regeneration response. As the population of pigment cells increases, the differentiation

step from progenitor cells to mature pigment cells is slowed down, and progenitor cells are

accumulated again. These explanations corroborate with the observations of the depletion of

punctate cells during depigmentation, and the much larger population of punctate cells than

pigment cells. Second, ans positive cells may also contribute to the differentiation of other

related cell types. During homeostasis, progenitor cells adopt a balanced contribution toward

pigment cells and other cell types (Figure 15A). In the event of depigmentation, contribution

toward pigment cells is likely prioritized over other cell types (Figure 15B). These explanations

corroborate with the observation of extensive co-expression between punctate and pigment

markers during early repigmentation.

4.2 Regulation of depigmentation and pigment cell regeneration

A number of genes were screened by RNAi to investigate their functions in depigmentation and

pigment cell regeneration. Among the genes with down-regulation in depigmented animals,

epimerase, hmbs and alas were observed to regulate depigmentation. Among both the up- and

down-regulated genes, however, regulators of pigment cell lineage specification were not

identified. Nevertheless, RNAi studies of draper and fst, which are candidate genes with reported

functions in cell corpses removal and missing tissue response, generated intriguing phenotypes

(Zhou et al., 2001; Freeman et al., 2003; Gaviño et al., 2013). The RNAi screens proved the

principle that important regulators of pigment cell biology can be identified by analysis of

differentially expressed genes, especially for genes with roles in depigmentation. It is noticed,

however, that essentially no transcription factors were picked up in the RNAseq data with a two-

fold change cut-off in differential expression of depigmented animals. It is possible that some

transcription factors only require subtle changes in their own expression to initiate transcription

of the downstream target and amplify the effect of its transcriptional regulation. These subtle

changes are likely masked by the overwhelming background of transcriptional events in

unrelated cell types in the whole worm sample used for RNAseq. Considering the relatively large

sizes of animal samples used for RNAseq in this study, the minute changes of transcription


factors in a small portion of pigment and pigment progenitor cells are likely not picked up by the

whole worms RNAseq.

Demonstrated by the hmbs RNAi experiment, the presence of pigment is required for the light-

induced pigment cell loss (Figure 12B). This observation supports the hypothesis that planarian

pigment generates ROS after light exposure and thus lead to pigment cell death. More

importantly, this finding demonstrates that light exposure induced cell ablation is specific to

pigment-containing cells. It indicates that the novel method of light-induced depigmentation

would indeed allow us to identify regulators specific to the pigment cell lineage. Also, it means

that pigment progenitor cells are unlikely to be ablated by light exposure, due to their lack of

pigment. The depletion of pigment progenitor cells is likely due to the rapid pigment cell

maturation and rate-limiting ASC differentiation, as proposed earlier (Figure 15A, B).

The decreased expression level of epimerase in depigmented animals (from RNAseq results)

suggests that epimerase expressing cell may sacrifice themselves during the potential engulfment

and gut excretion of cellular corpses. The gut expression pattern of epimerase corroborates with

its potential gut excretion function during depigmentation. It is unlikely for all types of cells

undergone gut excretion, including those induced by p53 RNAi or mex-3 RNAi, to be capable of

migration to gut for clearance. In comparison, a commonly used mechanism, such as engulfment

and transportation by macrophage-like cells, is more likely to exist. If the macrophage-like cells

themselves are lost during the process of gut excretion, then it explains the observation of a

reduced expression in depigmented animals, a state that requires intensive gut excretion and

possibly major loss of these cells. It is possible that the epimerase identified here is enriched in

such macrophage-like cells and plays a crucial role in the engulfment and gut excretion processes

(Figure 16).

Two possible functions of draper may account for its involvement in depigmentation. First,

draper may function in planarians similar to its homolog in other organisms by regulating cell

corpse removal (Zhou et al., 2001; Freeman et al., 2003). Thus, its functional perturbation leads

to reduced pigment cell removal via gut excretion (Figure 16). Second, draper may regulate the

location and the rate of proliferation of multiple cell types, including pigment cells, as suggested

by the ectopic piwi-1 and prog-2 expressions in draper RNAi (Figure 16). I was not able to

quantify the density of pigment cells under draper RNAi condition due to the altered cellular


morphology and the diminished cellular boundaries. But if pigment cells are indeed over-

proliferated during draper knockdown, then it may explain the increased residual population of

pigment cells during light exposure.

I tested the functional involvement of fst in pigment cell regeneration. I hypothesized that as a

positive regulator of missing tissue regeneration response, fst should not affect pigment cell

regeneration during repigmentation due to a lack of physical injury. Indeed, fst RNAi did not

reduce the rate of pigment cell regeneration. But on the contrary, fst increased the rate of pigment

cell proliferation during not only regeneration but also homeostasis. In comparison to missing

tissue response, pigment cell proliferation is likely positively regulated by the TGF-𝛽 pathway.

And RNAi knockdown of fst, a bioneutralizer of the positive ligands including activin, naturally

disinhibits this pathway and leads to increased proliferation rate (Figure 16).

Figure 16 Model of epimerase, draper and fst functions in pigment cell lineage specification and excretion. epimerase positively regulates pigment cell death in response to light exposure and their removal by gut excretion. draper may be directly involved in the process of recognition and engulfment of dying pigment cells. draper may also be responsible for the maintenance of the compartmentalized lineage marker expression, negatively regulating ASC and progenitor differentiation to prevent ectopic expressions of these genes. fst negatively regulates the rate of pigment cell lineage specification during both homeostasis and repigmentation.


4.3 Future directions

4.3.1 Deeper analysis of pigment cell and potential progenitors

Further evidence can be gathered to support ans positive cells as pigment cell progenitors. If

punctate cells marked by ans are indeed progenitors, then a higher percentage of PIWI protein

co-expression shall be observed in punctate cells than in mature pigment cells. The piwi

transcripts are well-established markers for the ASC population in planarians (Reddien et al.,

2005). The PIWI protein, on the other hand, persists slightly longer than the piwi transcripts in

cells that have recently undergone differentiation. It is thus a good marker of early progenies. A

higher percentage of PIWI co-expression in punctate cells would indicate that punctate cells are

more upstream in the lineage specification than the mature pigment cells. Similarly, BrdU, a

thymidine analog, can be used to detect recent DNA replications (Newmark and Sánchez

Alvarado, 2000). In planarians, ASCs are the only dividing cells (Eisenhoffer et al., 2008). The

rate of BrdU signal appearance in the post-mitotic tissue is a direct indicator of their turnover

rate from ASCs. If BrdU is sooner and more abundantly detected in punctate cells than mature

pigment cells, then punctate cells are more upstream in the lineage specification.

Earlier I presented a model that pigment cell lineage specification may pause at punctate cell

stage, and these cells act as a reserve for immediate pigment cell regeneration response. To test

this model, the regeneration of pigment cells in late-repigmenting and lethally irradiated (using 𝛾

irradiation to eliminate all dividing cells) animals can be analyzed. It has been demonstrated that

at later stage of repigmentation, the population of punctate cells is much greater than that of

pigment cells (Figure 11A, B). At the same time, the regeneration of pigment cells is incomplete.

If the pigment cells can continue to regenerate after irradiation, without additional ASC

divisions, then it would strongly argue punctate cells as an intermediate stage of lineage

specification, and as progenitors of mature pigment cells.

4.3.2 Continued screening of pigment cell lineage markers and regulators

In order to overcome the low detection rate of differentially expressed transcription factors in the

current RNAseq data of depigmentation time course, a new RNAseq dataset was generated.

Whole worm samples much smaller in size were used to produce the new RNAseq dataset. One

finding in this new RNAseq dataset is that the pigment cell marker hmbs established previously


shows the highest level of down-regulation in the depigmented samples, with a striking 109-fold

reduction in the transcript level, comparing to that of 13-fold in the old RNAseq data. With the

smaller worm samples, changes in the small subset of pigment cell population can be more

readily detected in the whole worm RNAseq, especially for the subtly changing transcription

factors. Using a hidden Markov model, a list of potential transcription factors has also been

generated based on homology to known transcription factors in other organisms and conserved

DNA binding domains. More pigment cell lineage markers and regulators should be screened

using a combination of these new datasets.

In addition to the whole worm RNAseq datasets, I have also attempted isolation of pigment cells

for single-cell RNAseq by fluorescent activated cell sorting (FACS). No fluorescent stains were

used for labeling of pigment cells during the past attempts. It is important to continuing

optimizing the detection of absorption by pigment cells in various wavelength. Once the

absorption profile of pigment cells is established by comparing cell suspensions prepared from

pigmented and unpigmented animals, the pigment cells can then be isolated by the cell sorter

based on these parameters. It is also shown in a recently published study that single cell

transcriptomes can be generated from RNAseq of de-cross-linked cellular samples (Thomsen et

al., 2015). Robust fluorescent labeling of pigment cells by hmbs has been generated by dFISH

with a formaldehyde fixation step. Once the de-cross-link method is optimized in the planarian

system, fixed cells sorted by FACS with hmbs marker can then be used for generation of pigment

cell transcriptome. Specific transcription factors expressed and required for pigment cell identity

maintenance can then be identified in this transcriptome.

A recent report has established subclasses within the planarian stem cell compartment, with

different subclasses contributing to different lineages of tissue types (van Wolfswinkel et al.,

2014). It is crucial to test if the transcription factors that determine the epidermal/ectoderm

lineage, such as zfp-1¸ can specifically regulate the regeneration of pigment cells. The effect of

zfp-1 on pigment cell regeneration can be compared to that of hnf4 and gata4/5/6

(endoderm/intestinal lineage) to determine which lineage are pigment cells more related to (van

Wolfswinkel et al., 2014). RNAi-based functional analysis of these transcription factors during

planarian repigmentation would allow us to at least categorize pigment cells into the general

subclasses of the tissue lineages they define.


4.3.3 Further analysis of depigmentation regulators

The level of ROS under epimerase and draper RNAi conditions can be detected by published

methods, including localization of ROS in the planarian tissue (Tarpey et al., 2004). This is key

to determining whether these genes function to reduce the ROS level, or to excrete dying

pigment cells during light exposure. The expression patterns of epimerase and draper should

also be generated in depigmented animals, in order to determine if they have dynamic expression

patterns during depigmentation and gut excretion. These experiments will be able to associate

epimerase and draper with their potential functions. In addition, the level of apoptosis in

normally pigmented and depigmenting animals should be determined using TUNEL as

previously described (Lin and Pearson, 2014). An increased level of TUNEL stains is expected

in epimerase or draper RNAi animals if these genes indeed involved in the engulfment of dying

cells and the gut excretion. The knockdown of these genes will lead to an accumulation of dying

cells that can be detected by TUNEL.

In order to establish epimerase and draper as regulators for gut excretion, it is key to

demonstrating that they regulate other types of gut excretion as well. These include the mex-3

RNAi induced prog-2 positive cell excretion, and the p53 RNAi induced cycB positive cell

excretion (Zhu et al., 2015; Pearson and Sánchez Alvarado, 2010). mex-3 and p53 RNAi

treatments should be performed in epimerase or draper RNAi background to determine if either

gut excretion process is affected.

Further investigation is necessary to better characterize the mechanisms leading to the ectopic

expressions of piwi-1 and prog-2 observed in draper RNAi animals. Phospho-histone H3 (H3P)

is a specific marker of dividing cells during metaphase. H3P signal and BrdU incorporation

studies can be performed in draper RNAi worms to determine if the cells that are ectopically

expressing piwi are also actively dividing. This experiment will provide evidence on whether

these cells contribute to ectopic tissue differentiation and outgrowth.

4.3.4 Further analysis of pigment cell regeneration regulators

fst is the only regulator of pigment cell regeneration investigated in this thesis. For fst, and for all

future pigment cell regeneration regulators to be discovered, the specificity of its regulation in

the pigment cell lineage should be determined. The proliferation of multiple tissue types under


RNAi conditions of various regulators can be determined by monitoring the level of BrdU

incorporation and the extent of PIWI protein co-expression in relation to expressions of lineage

specific markers, such as gata4/5/6, chat, hmbs, and ans (van Wolfswinkel et al., 2014; Currie

and Pearson, 2013). For a specific regulator of pigment cell lineage, it is expected that the

pigment cell proliferation shows the most dramatic changes under RNAi conditions of the

regulators while the other lineages are largely unaffected. Otherwise, the regulators would be

deemed to function in a wider range of differentiation activities and are not specific to the

pigment cell lineage.

4.3.5 Comparative transcriptomics for identification of gut excretion regulators

It is likely that active regulators of planarian gut excretion will display altered expression level

during gut excretion events. Looking at only one type of gut excretion event, however, is hard to

differentiate between true regulators of gut excretion from other irrelevant genes with altered

expression levels. For instance, wound response genes, including egr-3, hadrian and smed-mapk,

are up-regulated in depigmented animals, potentially confound the identification of regulators

specific for gut excretion (Wenemoser and Reddien, 2010; Wenemoser et al., 2012). On the

other hand, comparative transcriptomics by analyzing commonly altered gene expressions

among different gut excretion events would allow identification of key shared regulators. Genes

identified with this method can then be tested in each of the gut excretion events, including light

exposure-induced, mex-3 RNAi induced, and p53 RNAi induced to verify their functions.



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Appendix 1: List of in situ patterns identified but not included in the main text


Appendix 2: List of genes cloned and screened in this study

Transcript Name Up/Down Primer Sequence Gene/Homolog in situ? RNAi?






















COE1 Yes Yes









































Transcript Name Up/Down Primer Sequence Gene/Homolog in situ? RNAi?
































































Transcript Name Up/Down Primer Sequence Gene/Homolog in situ? RNAi?
































































Transcript Name Up/Down Primer Sequence Gene/Homolog in situ? RNAi?

































Appendix 3: List of phenotypes observed during RNAi screen

Clone # Transcript Expression in

Depig animals Gene Name Phenotype

pXW0031 A002843 Down 13.5X hmbs depigmentation, enlarged body size

pXW0046 A011154 Down 5.0X epimerase impaired Light Induced Depigmentation (LID)

pXW0047 A012985 Down 4.9X alas depigmentation, death

pXW0073 A058936 Up 2.9X N/A diffused eyes during LID

pXW0080 zfp-1 death

pXW0107 A002380 Down 2.9X Polyubquitin-C reduced LID

pXW0109 A004241 Down 29.8X olfactory receptor 493 reduced LID

pXW0115 A016646 Down 17.8X transcription factor COE1 reduced LID

pXW0113 A015989 Down 2.4X (Cation exchanger 5) reduced body size

pXW0119 A017725 - ATR death

pXW0120 A006481 Down 1.3X draper wrinkle skin, humps (out growth), reduced LID

pXW0121 A011934 Down 2.6X lmo fragment reduced LID, increased rate of pigment cell regeneration

pXW0122 A012033 Down 2.4X shoc-2 reduced LID

N/A A012581 Down 1.1X follistatin, fst Impaired blastema formation, increased rate of pigment cell

homeostasis and regeneration