Nuclear inheritance and genetic exchange in Giardia intestinalis, a divergent eukaryote with two nuclei

by

Meredith Lauren Carpenter

A dissertation submitted in partial satisfaction of the

requirements for the degree of

Doctor of Philosophy

in

Molecular and Cell Biology

in the

Graduate Division

of the

University of California, Berkeley

Committee in charge:

Professor W. Zacheus Cande, Chair Professor Kathleen Collins Professor Michael Eisen Professor Arash Komeili

Spring 2011

Nuclear inheritance and genetic exchange in Giardia intestinalis, a divergent eukaryote with two nuclei

© 2011

by Meredith Lauren Carpenter Abstract

Nuclear inheritance and genetic exchange in Giardia intestinalis, a divergent eukaryote with two nuclei

by

Meredith Lauren Carpenter

Doctor of Philosophy in Molecular and Cell Biology

University of California, Berkeley

Professor W. Zacheus Cande, Chair

The divergent eukaryotic parasite Giardia intestinalis is a major cause of diarrheal disease worldwide. The Giardia life cycle consists of two major stages: the binucleate trophozoite, which is found attached to the wall of the small intestine, and the infectious cyst, which is able to persist for weeks in the environment. The two nuclei in the trophozoite are equivalent and remain independent during mitosis. Although Giardia is presumed to be asexual based on the lack of an observed sexual cycle or gametes, its genome contains low levels of heterozygosity. This identity between nuclei is surprising because an asexual organism with two nuclei would be expected to accumulate differences due to genetic drift. Interestingly, the Giardia genome also contains seven homologs of meiosis- specific genes (HMGs): Spo11, Hop1, Dmc1a, Dmc1b, Hop2, Mnd1, and Mer3. In sexual organisms, these genes are specifically required for meiotic homologous recombination. Is Giardia using these genes for a cryptic sexual or parasexual process? Or, as an early-diverging eukaryote, is it using them for a process such as DNA damage repair, which is thought to have been the predecessor of meiotic homologous recombination? Furthermore, how is Giardia able to maintain two independent nuclei with low levels of heterozygosity? In this dissertation, I describe my efforts during my graduate studies to answer these questions. To do so, I set out to examine the functions of the HMGs in Giardia and to determine whether genetic exchange occurs between nuclei at some point in the Giardia life cycle. Before I could test the functions of the HMGs, I needed molecular tools for gene knockdown in Giardia. Attempts to knock down genes in Giardia using RNAi have not been successful, and knockouts are infeasible due the parasite’s tetraploid genome content. Thus, in Chapter 2, I describe my development of morpholinos as a new tool for gene knockdown in Giardia. When electroporated into trophozoites, these translation-blocking oligonucleotides are able to reduce the protein levels of their targets for up to 72 hours.

1 Next, I examined the behavior of the nuclei and cytoskeleton throughout the Giardia life cycle, with the goal of determining whether the quadrinucleate cyst is formed by a mitotic division or cell fusion and whether the nuclear pairs remain associated throughout encystation and excystation. In Chapter 3, I use cell mixing experiments and immunofluorescence staining of cytoskeletal components to demonstrate that cysts are formed by an incomplete mitotic division, and the parental nuclear pairs remain associated throughout encystation and excystation. Thus, any genetic exchange in the cyst occurs between nuclei derived from the same parent cell, and nuclear sorting does not appear to be a method by which Giardia reduces heterozygosity between nuclei. Finally, in Chapter 4, I demonstrate that nuclear fusion and genetic exchange occur between nuclei in the cyst, a process we have termed “diplomixis.” Although three of the HMGs (Dmc1b, Hop2, Mnd1) are expressed in the trophozoite, suggesting that they may play roles in somatic DNA damage repair/recombination, three HMGs (Spo11, Hop1, and Dmc1a) are only expressed in the cyst. Additionally, using integrated chromosomal markers, I have demonstrated that the nuclei fuse and exchange genetic material during encystation. Homologous recombination between nuclei, presumably catalyzed by the HMGs, could be the mechanism by which Giardia maintains low levels of allelic heterozygosity. These findings are significant not only for our knowledge of the basic biology and life cycle of this important parasite, but also for our understanding of the evolution of eukaryotic DNA damage repair and meiosis.

2 Table of Contents

ABSTRACT ...... 1 TABLE OF CONTENTS...... i LIST OF FIGURES AND TABLES ...... iii ACKNOWLEDGEMENTS ...... iv CHAPTER 1: AN INTRODUCTION TO GIARDIA INTESTINALIS AND THE EVOLUTION OF SEX AN INTRODUCTION TO GIARDIA INTESTINALIS ...... 1 Importance as a parasite...... 1 The Giardia life cycle...... 1 Trophozoite...... 1 Encystation...... 2 Excystation ...... 2 Genetic diversity in Giardia...... 3 Evolutionary position...... 3 GIARDIA AND THE EVOLUTION OF SEXUAL REPRODUCTION ...... 4 The evolution of sex ...... 4 Molecular signatures of sex...... 5 Sex in protists ...... 7 Giardia: asexual or just shy?...... 8 REFERENCES ...... 9 CHAPTER 2: USING MORPHOLINOS FOR GENE KNOCKDOWN IN GIARDIA INTESTINALIS ABSTRACT ...... 22 INTRODUCTION ...... 22 RESULTS ...... 23 Knockdown of GFP with a translation-blocking morpholino...... 23 Knockdown of GiKIN2b in wild-type cells causes extreme shortening of the flagella ...... 24 Using an epitope-tagged protein to measure morpholino knockdown ...... 25 DISCUSSION ...... 26 General considerations for morpholino use in Giardia ...... 26 Insights into kinesin-2 function in Giardia...... 27 Conclusions...... 28 MATERIALS AND METHODS ...... 28 REFERENCES ...... 31 CHAPTER 3: NUCLEAR BEHAVIOR AND THE CYTOSKELETON DURING DIFFERENTIATION OF GIARDIA INTESTINALIS ABSTRACT ...... 49 INTRODUCTION ...... 49 RESULTS ...... 51

i Cysts are formed by an incomplete mitotic division ...... 51 Generation of disc fragments in the cyst ...... 52 Reconstruction of the cytoskeleton during excystation...... 52 DISCUSSION ...... 53 MATERIALS AND METHODS ...... 55 REFERENCES ...... 57 CHAPTER 4: NUCLEAR FUSION AND GENETIC EXCHANGE BETWEEN NUCLEI IN THE GIARDIA INTESTINALIS CYST ABSTRACT ...... 71 INTRODUCTION ...... 71 RESULTS ...... 72 Three HMGs are expressed only during encystation...... 72 Genetic exchange occurs between nuclei in the cyst ...... 73 Cyst nuclei display dynamic movements...... 74 DISCUSSION ...... 74 MATERIALS AND METHODS ...... 77 REFERENCES ...... 79

ii List of Figures and Tables Chapter 1 FIGURE 1.1: The life cycle of Giardia intestinalis ...... 17 FIGURE 1.2: Phylogenetic tree of major eukaryotic supergroups and evidence for sexual reproduction ...... 19 TABLE 1.1: Phylogenetic distribution of meiosis-specific genes in eukaryotes and their prokaryotic homologs...... 21

Chapter 2 TABLE 2.1: Sequences of the morpholinos used in this study ...... 34 FIGURE 2.1: Time course of GFP knockdown by morpholinos...... 35 FIGURE 2.2: Illustration of gating parameters used in Figure 2.1...... 37 FIGURE 2.3: Representative histograms of flow cytometry data presented in Figure 2.1...... 39 FIGURE 2.4: Doubling the concentration of morpholinos does not result in a substantial increase in levels of knockdown...... 41 FIGURE 2.5: Efficiency of electroporation of morpholinos at different concentrations ...... 43 FIGURE 2.6: Translation-blocking morpholinos reduce GiKIN2b protein levels and cause extreme shortening of the flagella...... 45 FIGURE 2.7: GFP-tagged GiKIN2b is knocked down by the anti-GiKIN2b morpholino...... 47

Chapter 3 FIGURE 3.1: Overview of alternative theories for cyst formation and nuclear distribution during excystation...... 62 TABLE 3.1: Cysts are not formed by the fusion of two trophozoites...... 64 FIGURE 3.2: The Giardia cyst is formed by an incomplete mitotic division ...... 65 FIGURE 3.3: Development of the adhesive disk during encystation and excystation...... 67 FIGURE 3.4: Reorganization of the microtubule cytoskeleton during excystation...... 69

Chapter 4 FIGURE 4.1: Expression of the meiosis-specific gene homologs in trophozoites and cysts ...... 83 FIGURE 4.2: TUNEL staining increases during encystation ...... 85 FIGURE 4.3: Scheme for the creation and verification of a clonal strain with an integrated marker...... 87 FIGURE 4.4: Nuclear identity and genetic exchange during Giardia differentiation...... 89 FIGURE 4.5: Dynamic nuclear movements in the Giardia cyst...... 91 FIGURE 4.6: Beating of internal axonemes in the Giardia cyst ...... 93 FIGURE 4.7: Model of diplomixis ...... 95

iii Acknowledgements

This dissertation would not exist without the mentors, colleagues, friends, and family who have supported me at every step along the way. First and foremost, I would like to thank Zac and all the current and former members of the Cande lab for their encouragement, support, friendship, and mentorship. I could not have asked for better colleagues or friends. To Zac: Thank you for allowing me to follow my own path and pursue my own interests. Your tireless enthusiasm is infectious and often helped me persevere when things got tough. I am honored to have been your last graduate student. To my officemates Alex and Lil: Thanks for making work fun and always being willing to listen when I needed advice, a sounding board, or just an opportunity to vent. Your influence has definitely helped me to become a better scientist. To my parents: From buying me rocks for my rock collection to letting me raise fruit flies in our guest room, you have always supported me and my interest in science. Thank you for being the strongest believers in what I can accomplish, as well as my biggest cheerleaders. And last but certainly not least, to Greg: Your love and friendship means everything to me. I could never have made it this far without your support, and I’m excited to embark on the next phase of our life together. I love you.

iv Chapter 1

An introduction to Giardia intestinalis and the evolution of sex

An introduction to Giardia intestinalis “I have at times seen very prettily moving animalcules, some rather larger, others somewhat smaller than a blood corpuscle, and all of one and the same structure. Their bodies were somewhat longer than broad, and their belly, which was flattened, provided with several paws, with which they made such a movement through the clear medium…But although they made a rapid movement with their paws, yet they made but slow progress.” Antony van Leeuwenhoek, describing what were likely Giardia trophozoites, which he observed while looking at his own stool under a microscope (Dobell, 1920; Ford, 2005).

Importance as a parasite Giardia intestinalis (syn. Giardia lamblia, Giardia duodenalis) is a protozoan intestinal parasite that is a major cause of waterborne diarrheal disease worldwide (Savioli et al., 2006). Giardia was first described by Antony van Leeuwenhoek in 1681 when he examined his own watery excretions under an early microscope (Dobell, 1920; Ford, 2005). In the United States alone, an estimated 2 million cases of giardiasis occur each year (Mead et al., 1999); the disease is especially common in children and people who have traveled to developing countries (Yoder et al., 2010). The symptoms of giardiasis include diarrhea, abdominal cramps, bloating, weight loss, and malabsorption, although the disease can also be asymptomatic (Hellard et al., 2000; Thompson, 2000). The most commonly used drug for the treatment of Giardia infection are metronidazole, quinacrine, and albendazole; however, due to the spread of drug resistance, the search continues for alternative treatments (Wright et al., 2003). In addition to humans, other mammals can become infected with Giardia, including dogs, cats, domestic livestock, and even beavers (the source of the nickname for giardiasis, “beaver fever”). While some strains of Giardia are host-specific, some appear to have a broader host range and can be transmitted zoonotically (Thompson, 2000).

The Giardia life cycle Trophozoite Giardia is classified in the order Diplomonadida, whose members contain two nuclei. The Giardia life cycle consists of two main stages: a binucleate trophozoite stage and a quadrinucleate cyst stage (Figure 1.1 and (Adam, 2001)). When living attached to the surface of the small intestine, Giardia is found in the vegetatively dividing, flagellated trophozoite form. The trophozoite is a teardrop-shaped cell, and its two nuclei are both transcriptionally active, diploid, and contain complete copies of the genome (Bernander et al., 2001; Kabnick and Peattie, 1990; Yu et al., 2002). 1 The two nuclei remain separate during mitosis in the trophozoite, with each daughter cell inheriting two genetically distinct nuclei (Sagolla et al., 2006). The trophozoite can be grown in vitro in culture in medium containing (among other nutrients) bovine serum, high concentrations of cysteine, and bile salts (Keister, 1983). Giardia is microaerophilic, so it can be grown in closed tubes without the requirement of a completely anaerobic environment (Adam, 2001). One of the most prominent features of the Giardia trophozoite is its microtubule cytoskeleton (reviewed in (Dawson, 2010)), which includes the following components: four pairs of flagella (anterior, posterior, ventral, and caudal); a cluster of microtubules of unknown function called the “median body”; and the ventral adhesive disk, which is used by the cell to attach to substrates, likely via a suction- based mechanism (Figure 1.1) (Hansen et al., 2006). The ability of Giardia to attach to the intestinal epithelium allows it to remain in the upper and middle sections of the small intestine (duodenum and jejunum, respectively) and avoid being swept downstream by peristalsis (Adam, 2001).

Encystation If a trophozoite detaches from the intestinal wall and travels into the colon, it begins to encyst. Encystation is thought to be triggered by a combination of alkaline pH and cholesterol starvation, which can be mimicked in vitro by using medium with high concentrations of bile salts and a pH of 7.8 (Gillin et al., 1989; Gillin et al., 1988; Kane et al., 1991; Lujan et al., 1996; Reiner et al., 1995). The trophozoite has a 4N genetic content (2N = 10 in each nucleus), and its content increases to 16N (4 x 4N nuclei) in the mature cyst (Bernander et al., 2001). The trophozoite cytoskeleton is largely disassembled in the cyst, with internal axonemes and fragments of the adhesive disk the only prominent structures (Buchel et al., 1991; Buchel et al., 1987; Erlandsen et al., 1989; Hetsko et al., 1998). The hardy cyst is surrounded by a tough wall composed primarily of leucine- rich cyst wall proteins and a ß(1-3)-N-acetyl-D-galactopyranosamine homopolymer (Gerwig et al., 2002). During encystation, the components of the cyst wall are transported to the periphery of the cell in Golgi-like encystation-specific vesicles (Faubert et al., 1991; Stefanic et al., 2006). Encystation allows the cyst to survive a range of conditions, including exposure to water, until it reaches the small intestine of its next host. A dose as small as 10 cysts can establish infections in humans (a number determined in the 1950s by feeding “prisoner volunteers” capsules containing different numbers of Giardia cysts), and hosts shed thousands to millions of cysts per gram of stool (Danciger and Lopez, 1975; Rendtorff, 1954).

Excystation When a cyst is ingested by its next host, it must first pass intact through the acidic environment of the stomach before it can establish infection in the small intestine. In fact, this exposure to acid is the first trigger for excystation, the process by which the cell exits the cyst and transforms back into a trophozoite. This process can be recapitulated in vitro by a two-stage method in which cells are first

2 exposed to an acidic solution (pH 4.0) followed by a more basic solution containing proteases (Bingham and Meyer, 1979; Boucher and Gillin, 1990; Hautus et al., 1988; Isaac-Renton et al., 1986; Rice and Schaefer, 1981). The quadrinucleate excyzoite exits the cyst through one pole, flagella first, and quickly reorganizes and reassembles its cytoskeleton (Buchel et al., 1987; Coggins and Schaefer, 1984, 1986; Erlandsen et al., 1989). The cell eventually undergoes cytokinesis to produce two daughter trophozoites; because these cells are 8N (2 x 4N nuclei), they are thought to divide again immediately without the need for an intervening S phase (Bernander et al., 2001). This ability for one cyst to rapidly produce four daughter trophozoites has been proposed as a mechanism by which low numbers of Giardia cysts are able to produce infection (Bernander et al., 2001).

Genetic diversity in Giardia The species known as Giardia intestinalis actually consists of at least seven genotypes (called “assemblages”), of which only two, assemblages A and B, are known to infect humans (Adam, 2001). These are also the most well-studied assemblages in vitro, with the dominant laboratory strain WB representing assemblage A and the strain GS representing assemblage B. The genome sequences of both WB and GS have been determined, as has that of assemblage E isolate P15 (Franzen et al., 2009; Jerlstrom-Hultqvist et al., 2010; Morrison et al., 2007). The differences between these strains include insertion-deletion events, single- nucleotide polymorphisms, and chromosomal rearrangements; these differences, combined with overall nucleotide identities ranging from 70-80%, have led some to argue that these assemblages actually represent different species (Franzen et al., 2009; Jerlstrom-Hultqvist et al., 2010). Within populations, the levels of allelic heterozygosity are quite low, ranging from 0.5% in GS to less than 0.01% in WB and P15 (Franzen et al., 2009; Jerlstrom- Hultqvist et al., 2010; Morrison et al., 2007), and its populations are largely clonal (Tibayrenc, 1993; Tibayrenc and Ayala, 2002; Tibayrenc et al., 1990). The finding of such low levels of heterozygosity in a supposedly asexual organism with two nuclei was unexpected and has led to the hypothesis that Giardia undergoes some sort of sexual or parasexual cycle to help reduce heterozygosity between its two nuclei (see discussion below).

Evolutionary position Giardia is one of the most divergent eukaryotes studied to date and a member of what is often classified as one of the earliest-diverging eukaryotic lineages based on single- and multi-gene phylogenies (Arisue et al., 2005; Baldauf et al., 2000; Baldauf, 2003; Best et al., 2004; Ciccarelli et al., 2006; Morrison et al., 2007). In addition, it is anaerobic, with highly reduced mitochondria, no peroxisomes, and no recognizable Golgi apparatus in the trophozoite (Abodeely et al., 2009; Adam, 2001; Marti et al., 2003; Regoes et al., 2005). Giardia also has few introns in its ~11.7-Mb sequenced genome (Morrison et al., 2007). 3 Although eukaryotic rooting schemes remain controversial, in three of the most popular hypotheses, Giardia belongs to one of the most divergent eukaryotic groups from commonly studied organisms like animals and yeast (Figure 1.2) (Fritz- Laylin et al., 2010). It is most parsimonious to assume that features shared between Giardia and another major eukaryotic group likely existed in their common ancestor. If these groups diverged early in eukaryotic evolution, these shared features were likely present early in eukaryotic evolution. Likewise, features absent in Giardia, if also absent in its free-living relatives, could imply that these features involved later. Thus, comparative studies of Giardia biology have the power to shed light on characteristics of the last common ancestor of all eukaryotes.

Giardia and the evolution of sexual reproduction “In many organic beings, a cross between two individuals is an obvious necessity for each birth; in many others it occurs perhaps only at long intervals; but in none, I suspect, can self-fertilisation go on for perpetuity.” Charles Darwin, On the Origin of Species (Darwin, 1859).

The evolution of sex When, why, and how did sex evolve? These questions have intrigued scientists, including Darwin, for centuries; in fact, the existence of sex has been called “the queen of problems in biology” (Bell, 1982). Meiosis, the special reductional cell division that produces gametes for sexual reproduction, is ubiquitous among vertebrates and plants; however, a number of eukaryotes, including certain protists, are thought to reproduce only asexually. The existence of asexual eukaryotes has been a topic of intense interest because of its implications for our understanding of the benefits and disadvantages of sexual reproduction. Although the debate continues about why sex evolved, several popular theories have emerged. The main disadvantage of sex is referred to as the “twofold cost of sex,” a term first coined by John Maynard Smith (Maynard Smith, 1978) referring to the fact that in sexual populations with two sexes, only one sex is able to produce offspring. Thus, sexual populations have half the level of fitness should be outcompeted by asexual populations, which have the capacity to increase exponentially because all members are able to reproduce. Another disadvantage is that, in general, a sexually reproducing organism will only pass on half its genes to its progeny. Of course, many theories have also been put forward to explain the advantages of sex (one author counted 20 different theories in the early 1990s (Kondrashov, 1993)). The most popular theories can be briefly summarized as follows:

Creation of novel genotypes (combining advantageous alleles): Originally described by Fisher (Fisher, 1930) and Muller (Muller, 1932). In a population without recombination, if two beneficial mutations occur in different individuals, they can never be combined in a single descendent.

4 Instead, combination of these alleles would require that the mutations arise independently in a single individual.

Muller’s ratchet (removal of deleterious alleles) (Muller, 1964): Asexual populations are prone to accumulating mildly deleterious mutations due to the random loss of clonal lines without those mutations, ultimately leading to genome deterioration. The related deterministic mutation hypothesis (Kondrashov, 1988) posits that sexual recombination shuffles deleterious mutations such that some individuals inherit more mutations than others. These individuals with more mutations are likely to be less fit and thus die out, purging the mutations from the population.

Red Queen hypothesis (host-parasite coevolution) (Bell, 1982; Hamilton et al., 1990; Van Valen, 1973): In the evolutionary war between hosts and their parasites, sex allows populations to retain a reservoir of variation that may be advantageous under different conditions.

DNA repair and complementation (Bernstein et al., 1985; Bernstein et al., 1987): Homologous recombination during meiosis allows for the repair of DNA damage in the genome before it is transmitted to offspring; genetic variation is a byproduct of this process. Diploidy, initially transient, became obligate in many organisms as a way to compensate for recessive deleterious mutations.

However, obligate sex is not necessary to reap many of these benefits of sexual reproduction: rare sex (5-10%) can effectively stop Muller’s ratchet and remove deleterious mutations from the genome, as well as increase levels of variation and fitness in a population [see (D'Souza and Michiels, 2010) and references therein]. Additionally, from a parasite’s point of view, rare sex would allow the individuals best adapted to a certain host to produce highly fit clonal populations, but would also provide the option to vary in response to changing environments and host adaptations (Flatt et al., 2001).

Molecular signatures of sex As Muller put it, “the essence of sexuality…is recombination” (Muller, 1932). Accordingly, one line of evidence that has been used to conclude that sex was present in the last common ancestor of all eukaryotes is the presence of homologs of meiosis-specific genes (HMGs) in the genomes of members of all major eukaryotic groups (Malik et al., 2008) (Table 1.1). These genes are the core machinery of meiotic homologous recombination, and would be expected to be lost in obligate asexual organisms. They include the following main players (Malik et al., 2008):

Spo11: Type II topoisomerase that makes double-strand breaks (DSBs) in DNA to initiate homologous recombination during meiosis.

5

Hop1: Component of the synaptonemal complex that binds DSBs and oligomerizes during meiotic prophase I.

Hop2 & Mnd1: Form a complex and ensure accurate homology searching during meiotic prophase I. These proteins are required for stable heteroduplex formation and interhomolog repair.

Dmc1: RecA-family protein that assembles on resected single-strand DNA at DSBs and promotes strand invasion and favors interhomolog recombination (as opposed to its homolog Rad51, which favors repair of the sister chromatid).

Msh4 & Msh5: Form a heterodimer that interacts with the Mlh1/Mlh3 heterodimer and directs Holliday junction resolution towards crossover with interference.

Rec8: Meiotic homolog of the cohesin Rad21; maintains sister chromatid cohesion during meiosis I.

Mer3: Meiosis-specific DEAD-box helicase. Promotes Holliday junction resolution with crossover interference with the ZMM proteins, including Msh4 and Msh5.

However, the mere presence of these HMGs does not prove that the organism is using them for meiosis. Indeed, it has been proposed that these genes had a non- meiotic function in early eukaryotes, most likely in DNA damage repair (Marcon and Moens, 2005; Villeneuve and Hillers, 2001). In support of this hypothesis, the bacterial and archaeal homologs of several HMGs (including topoisomerase VI, MutS, MutL, and RecA) are used for DNA damage repair. Furthermore, asexual bdelloid rotifers have retained their HMGs (J. Logsdon, personal communication), and it has been recently shown that the fungus Candida albicans uses Spo11 for homologous recombination in a non-meiotic parasexual stage of its life cycle (Forche et al., 2008). Other molecular signatures of sex have been proposed, although these are largely hypothetical due to the dearth of confirmed obligately asexual organisms. One of the most intuitive is the prediction that in an asexual diploid organism, homologous chromosomes should diverge in sequence over time because the pressure to remain similar for proper synapsis during meiosis no longer exists. This divergence was dubbed the “Meselson effect” due to what appeared to be confirmation of this prediction by Matthew Meselson’s work on bdelloid rotifers (Mark Welch and Meselson, 2000); however, subsequent work has suggested that these rotifers are actually degenerate tetraploids (Mark Welch et al., 2008), calling into question the relevance of the Meselson effect in actual asexual populations.

6 Furthermore, a recent study of asexual Daphnia populations found evidence of widespread ameiotic recombination that reduced allelic heterozygosity at a faster rate than new mutations were introduced, again suggesting that neutral allelic divergence may have a limited utility as an indicator of ancient asexuality (Omilian et al., 2006). Another predicted signature of asexual populations is the loss of transposable elements (TEs). In sexual populations, deleterious TEs can spread and persist at some level in the population without harmful effects (Arkhipova and Meselson, 2000; Dolgin and Charlesworth, 2006; Schurko et al., 2009). However, asexual populations, while lacking the means by which TEs can spread, also lack the ability to clear these elements; therefore, it has been proposed that asexual lineages should eventually lose most or all of their TEs unless they can be “domesticated” to serve the host organism in some way (Arkhipova and Meselson, 2000; Dolgin and Charlesworth, 2006; Schurko et al., 2009). This loss of TEs has been confirmed in bdelloid rotifers, which lack many common retrotransposons found in other animal groups (Arkhipova and Meselson, 2000; Gladyshev and Arkhipova, 2010). Finally, in the absence of meiotic recombination, allele frequencies in asexual populations should demonstrate high levels of linkage disequilibrium and even complete linkage of markers over the entire genome (Schurko et al., 2009).

Sex in protists While it is evident that there was a loss of sex in asexual metazoan and plant lineages (i.e., the last common ancestor of these groups had the capacity for sexual reproduction), it remains unclear whether the last common ancestor of all eukaryotes was sexual. Investigating the presence or absence of sex in other major groups of eukaryotes has the potential to shed light on how sexual reproduction and the machinery for meiotic homologous recombination evolved. Despite the finding of HMGs in all protistan genomes sequenced thus far, of the major lineages of microbial eukaryotes, sexual reproduction has only been described in the Amoebozoans and Stramenophile-Alveolate-Rhizaria supergroup (Figure 1.2) (Malik et al., 2008). Few molecular studies of homologous recombination have been performed in these organisms; however, at least in Tetrahymena, the HMGs appear to play largely similar roles to those in eukaryotes like yeast, plants, and humans, mediating homologous recombination during meiotic prophase I (Loidl and Mochizuki, 2009; Loidl and Scherthan, 2004; Lukaszewicz et al., 2010; Mochizuki et al., 2008). Interestingly, there is also increasing (indirect) evidence for sexual reproduction in the Euglenozoans, namely Trypanosomes and Leishmania. Several recent studies have identified cryptic sexual stages of these organisms that take place in the tsetse fly/sandfly vectors and result in the production of recombinant lineages (Akopyants et al., 2009; Gaunt et al., 2003; Peacock et al., 2011). However, because the meiotic division in these organisms has not yet been characterized, the roles of the HMGs also remain unexplored, although it has been shown that Hop1,

7 Mnd1, and Dmc1 are expressed in the meiotic life cycle stage of T. brucei (Peacock et al., 2011).

Giardia: asexual or just shy? Giardia is presumed to be asexual based on the clonal structures of wild populations and the lack of observable mating types or gametes (Adam, 2001; Tibayrenc, 1993; Tibayrenc and Ayala, 2002; Tibayrenc et al., 1990). The genome also contains few TEs: retrotransposons have been identified, but they are located in subtelomeric regions and may have been co-opted for telomere maintenance (Arkhipova and Morrison, 2001; Burke et al., 2002; Prabhu et al., 2007; Ullu et al., 2005). However, as noted above, the Giardia genome contains very low levels of heterozygosity. Because an asexual organism with two independent nuclei would be expected to accumulate differences between the nuclei simply due to genetic drift, the finding of low levels of allelic heterozygosity in Giardia is surprising and has led to the suggestion that this organism is not, in fact, asexual (Birky, 2010). Consistent with this hypothesis, the Giardia genome contains at least seven HMGs (Malik et al., 2008; Ramesh et al., 2005): Hop1, Hop2, Mnd1, Dmc1a, Dmc1b, Spo11, and Mer3; it is missing homologs of Msh4, Msh5, and Rec8. Similarly, nine HMGs were found in Trichomonas vaginalis, which belongs to the same major eukaryotic group as Giardia. Additionally, a study of wild Giardia populations has revealed evidence of recombination within assemblage A2, again calling into question the presumed asexuality of the parasite (Cooper et al., 2007; Lasek- Nesselquist et al., 2009). Due to the evolutionary position of Giardia, determining whether it is indeed asexual or has a cryptic sexual cycle has implications for our understanding of how sex evolved during eukaryotic evolution. Furthermore, better knowledge of the life cycle of this parasite will impact our view of the evolution and spread of drug resistance in Giardia populations and could potentially aid in the development of treatments for giardiasis. The goals of the present study were threefold: 1) To understand the patterns of nuclear inheritance during the Giardia life cycle; 2) To determine whether Giardia undergoes a sexual or parasexual cycle in order to reduce heterozygosity between its nuclei; and 3) To establish whether the HMGs function in this process, or whether they have a non-meiotic function (for example, in DNA damage repair). To address these questions, I first sought to develop better tools with which to investigate gene function in Giardia. In Chapter 2, I describe the development of antisense morpholino oligonucleotides as a tool for gene knockdown in Giardia; morpholinos have now been used successfully in several recent studies (Carpenter and Cande, 2009; Garlapati et al., 2011; Paredez et al., 2011). In Chapter 3, I set out to examine in detail how Giardia’s two nuclei are inherited during encystation and excystation. This information about the parasite’s life cycle is crucial for our understanding of both its ability to maintain two identical nuclei and the patterns of genetic diversity in Giardia populations. Finally, in Chapter 4, I describe the discovery of “diplomixis”, a novel parasexual process in Giardia involving nuclear

8 fusion and genetic exchange, possibly facilitated by the HMGs, that takes place during encystation.

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16 Figure 1.1: The life cycle of Giardia intestinalis. The host ingests a cyst (4 x 4N nuclei = 16N overall), which passes through the stomach and subsequently excysts, dividing twice to form four 4N trophozoites. The trophozoite (2 x 2N = 4N in G1, 2 x 4N = 8N in G2) attaches to the wall of the small intestine and divides by mitosis. If a trophozoite is swept into the lower small intestine and then the large intestine, it encysts, reforming the 16N cyst that is eventually excreted from the body. Major cytoskeletal components of the trophozoite cell are as follows [top image]: the ventral adhesive disc (vd), used for attachment to the intestinal epithelium; four sets of flagella (afl = anterolateral; pfl = posterolateral; vfl = ventral; cfl = caudal); the median body (mb), a microtubule array of unknown function; and the two nuclei (n).

17 N N vd

mb afl afl

vfl pfl pfl

cflcfl 4N/8N trophozoite

Mitosis

Intestinal epithelium

16N cyst

Encystation Excystation

Excreted by host Persists in water supply Ingested by next host

18 Figure 1.2: Phylogenetic tree of major eukaryotic supergroups and evidence for sexual reproduction. Consensus cladogram of selected eukaryotes, highlighting five major groups with widespread support in diverse molecular phylogenies [adapted from (Fritz-Laylin et al., 2010; Parfrey et al., 2010)]. The polytomy indicates uncertainty regarding the order of early branching events. Representative taxa are shown on the right. The presence of meiosis and/or a sexual cycle in each lineage is represented by “+”; genetic evidence for sex is represented by “g”; and lineages in which there is no evidence for sex are indicated by “?”. The inset depicts three contending hypotheses for the eukaryotic root. Root 1: early divergence of unikonts and bikonts (Stechmann and Cavalier-Smith, 2002). Root 2: POD and JEH (together known as the “excavates”) branching first (Rodriguez-Ezpeleta et al., 2007; Yoon et al., 2008). Root 3: POD diverging first (Arisue et al., 2005; Ciccarelli et al., 2006; Morrison et al., 2007). Regardless of the rooting scheme, comparative studies involving Giardia have the potential to shed light on characteristics of the last eukaryotic common ancestor.

19 Eukaryotic supergroup Example Sex? Animals Homo Opisthokonts + Fungi Saccharomyces +

Dictyostelium Amoebozoans + Entamoeba ? Plants Zea Plantae + Green algae Chlamydomonas + Stramenophiles Thalassiosira + Rhizaria Gromia SAR + Apicomplexans Plasmodium Alveolates + Ciliates Tetrahymena + Heteroloboseans Naegleria ? Jakobids JEH Reclinomonas ? Euglenozoans Trypanosoma g Excavates Parabasalids Trichomonas ? POD Oxymonads Monocercomonoides ? Diplomonads Giardia g Eukaryotic rooting schemes unikont O O O A A A 1 P P P 2 S S bikont S 3 J J J P P P 20 Table 1.1: Phylogenetic distribution of meiosis-specific genes in eukaryotes and their prokaryotic homologs.

Organism Spo11 Hop1 Hop2 Mnd1 Dmc1 Msh4 Msh5 Rec8 Mer3

Bacteria - - - - RecA - - MutS Archaea TopoVI - - - RadA - Ski2

Giardia (G) + + + + ++ - - - +

Trichomonas + + ++ + + + + - +

Trypanosoma (G) + + + + + + + - +

Plasmodium (M) ++ + + + + - - - -

Dictyostelium (M) - ? + + ? + + ? ?

Arabidopsis (M) ++++ + + + + + + + +

Saccharomyces (M) + + + + + + + + +

Homo (M) + + + + + + + + +

The presence and number of homologs of each meiosis-specific gene are indicated by (+) and (-). The absence of a homolog from an unfinished or incompletely annotated genome is indicated by (?). (G) Genetic evidence for recombination. (M) Confirmed meiosis. Homologs present in the Giardia genome are highlighted in red. Table adapted from (Malik et al., 2007) and (Malik et al., 2008).

21 Chapter 2

Using morpholinos for gene knockdown in Giardia intestinalis

Abstract A major roadblock to studying Giardia biology is the dearth of techniques for gene knockdown. In this study, we used translation-blocking morpholinos to reduce protein levels in Giardia intestinalis. Twenty-four hours after electroporation with morpholinos targeting either green fluorescent protein (GFP) or the endogenous protein kinesin-2b, levels of these proteins were reduced by 60%. Kinesin-2b knockdown also produced cells with dramatically shortened flagella and missing or reduced median bodies. This phenotype was more extreme than the previously reported phenotype of a dominant-negative mutant of kinesin-2a, a kinesin-2b homolog likely found in the same complex. The kinesin-2b knockdown phenotype is consistent with a role for this protein, like kinesin-2a, in anterograde intraflagellar transport. We also found that levels of a GFP-tagged kinesin-2b are reduced by morpholinos to the same extent as the endogenous protein, although the knockdown occurred with slightly slower kinetics. Thus, an epitope-tagged transgene can be used as a reporter of morpholino efficacy for targets lacking specific antibodies. Due to their rapid action, ease of use, and stability in cell culture, we expect morpholinos to become a valuable new tool for characterizing gene function in Giardia.

Introduction Giardia intestinalis is a parasitic protist and a major cause of diarrheal disease in developing countries (Adam, 2001; Savioli et al., 2006). Giardia also belongs to what is perhaps one of the earliest diverging eukaryotic lineages (Morrison et al., 2007). For these reasons, this organism has been studied for years, both in the quest for new drug treatments and to shed light on the evolution of eukaryotic cells. However, certain aspects of giardial biology have proved intractable for researchers seeking to study gene function. The trophozoite contains two diploid nuclei, making the cell effectively tetraploid and gene knockouts by homologous gene replacement infeasible (Adam, 2001). In addition, although Giardia contains RNA interference genes (homologs of Dicer and Argonaute) and this machinery was recently implicated in the control of antigenic variation (Prucca et al., 2008; Saraiya and Wang, 2008), attempts to manipulate this system for gene knockdown have been unsuccessful (C.C. Wang, personal communication). While there have been many valuable tools developed for studying gene function in Giardia, a fast, reliable method to knock down genes is still lacking. A few researchers have used virus-mediated ribozyme constructs to achieve gene knockdown (Dan et al., 2000). However, drawbacks to this approach include the necessity to select for transformants, which may eliminate cells in which knockdown is deleterious, and the several days and high levels of drugs required for induction of the ribozyme. Additionally, in our experience, infection with giardiavirus can cause high levels of cell death. Due to these complications, the 22 ribozyme technique has not been adopted by the Giardia research community at large. Dominant-negative mutants are also used to study gene function (Dawson, 2007; Hoeng et al., 2008), but few genes are amenable to this approach. Another, more widely used approach has been to express the antisense sequence of the entire open reading frame of the target gene from a plasmid using a strong promoter (Lauwaet et al., 2007; Prucca et al., 2008; Touz et al., 2002). Presumably, these antisense sequences bind to their cognate transcript, producing long (>1000 bp) double-stranded RNAs. However, whether the subsequent reduction in RNA and protein levels is caused by a yet uncharacterized RNAi-mediated pathway that accepts extremely long dsRNA substrates, or whether it works solely by sequestering the mRNA from translation and prompting its destruction by cellular RNases, remains to be determined. Varying levels of knockdown (from 34- 100%) have been achieved by expressing the antisense sequence of large portions of the open reading frame of the target gene under a strong promoter (Gaechter et al., 2008; Lauwaet et al., 2007; Marti et al., 2003; Prucca et al., 2008; Touz et al., 2002). But because a promoter allowing for tight control of gene expression has not been developed for Giardia, this approach can only be applied to study non-essential (Prucca et al., 2008) or encystation-specific (Gaechter et al., 2008; Marti et al., 2003) genes. Furthermore, it is not possible to control for off-target effects when using this technique. In this study, we use morpholinos to knock down gene expression in Giardia. Morpholinos are modified antisense oligonucleotides in which a six-membered morpholine ring replaces the deoxyribose ring of DNA, and non-ionic phosphorodiamidate linkages replace the typical anionic phosphodiester linkages (Moulton and Yan, 2008). As a result, they cannot be degraded by cellular nucleases and are stable in cell culture (Summerton, 1999). When designed to bind between the 5’ cap and 25 nucleotides downstream of the translation start site of the target mRNA, morpholinos (typically 25-mers) will sterically block ribosome binding and prevent translation of the target protein (Summerton, 1999). These translation- blocking morpholinos, introduced by electroporation, have been used to prevent new protein synthesis in trypanosomes (Shi et al., 2007). Morpholinos can also be used to block splicing and render a target inactive; however, Giardia contains only four genes with introns, precluding the use of this strategy for most targets (Morrison et al., 2007). In the present study, we first chose to use a translation-blocking morpholino to target enhanced green fluorescent protein (eGFP, hereafter referred to as GFP) driven by a strong promoter, allowing us to test the effects of morpholinos on a neutral, exogenously derived gene expressed at high levels. Next, we targeted the endogenous Giardia intestinalis kinesin-2b (GiKIN2b), a paralog of the previously characterized kinesin-2a (Hoeng et al., 2008), and compared the resulting phenotype with the known phenotype of the GiKIN2a dominant negative.

Results Knockdown of GFP with a translation-blocking morpholino

23 To determine the efficacy of translation-blocking morpholinos in Giardia, we first targeted enhanced GFP in a strain expressing the gene under the glutamate dehydrogenase promoter (Yee et al., 2000). In this strain, diffuse GFP fluorescence is found throughout the cytoplasm (data not shown). The 25-mer morpholino was designed to target the first 24 bases of the GFP open reading frame, plus one base upstream of the start codon (Table 2.1). As a specificity control, we used a morpholino containing five mispaired bases (Table 2.1). The inclusion of five mispairs has been shown to destabilize pairing of the morpholino to its target unless the cytoplasmic morpholino concentration is extremely high; therefore, this control can act as a sensor for concentration-dependent off-target effects (Moulton and Yan, 2008). This control morpholino also shares its chemical properties and base composition with the experimental morpholino. After electroporation with either 10 µM or 100 µM morpholino, cells were grown for the indicated amount of time, then analyzed by flow cytometry (Figure 2.1). For flow cytometry, a gate for GFP-positive cells was created based on comparison to wild-type cells (Figure 2.2). In all cases, the presence of morpholino had no observable effect on cell growth compared to the no-morpholino control (data not shown). GFP levels in the no-morpholino and mispair controls remained approximately equal at all time points (Figure 2.1; see Figure 2.3 for representative flow cytometry histograms). However, twenty-four hours after electroporation, GFP levels in the cultures electroporated with 100 µM anti-GFP morpholino had decreased by ~60%, and remained at approximately this level for the next two days, only increasing after the cultures were passaged and allowed to grow for 24 hours (Figure 2.1, 96 hours). In the cultures electroporated with 10 µM anti-GFP morpholino, protein levels decreased by a maximum of ~40%. Treatment with a higher concentration of morpholino (200 µM) produced approximately the same level of knockdown as 100 µM (Figure 2.4). For both the GFP and GiKIN2b experiments described below, maximum knockdown was not achieved until 24 hours after electroporation (data not shown). This delay likely reflects the time needed for turnover of pre-existing protein in the cell, and/or dilution by cell division. To determine whether the remaining GFP-positive cells received morpholino, we electroporated cells with a fluorescently labeled anti-GFP morpholino. Twenty- four hours after electroporation, >99% of cells with 100 µM and 200 µM morpholino were positive for morpholino fluorescence, whereas only ~37% of cells with 10 µM morpholino were positive (Figure 2.5). However, within the morpholino-positive cells, there was no obvious correlation between morpholino fluorescence and GFP levels. The remaining GFP-positive cells in these experiments likely started out with more GFP (the population is heterogeneous due to variations in plasmid copy number), and did not receive enough morpholinos to reduce the GFP below the level of detection.

Knockdown of GiKIN2b in wild-type cells causes extreme shortening of the flagella

24 Next, we designed a morpholino to target an endogenous gene, GiKIN2b (Table 2.1). This gene is a homolog of the previously characterized giardial kinesin- 2a (GiKIN2a), which is involved in anterograde intraflagellar transport in Giardia (Hoeng et al., 2008). In the previous study, expression of a dominant-negative GiKIN2a mutant produced cells with shortened flagella (Hoeng et al., 2008). In many species, the kinesin-2 heterotrimeric complex includes two kinesin-2 homologues and a kinesin-associated protein (KAP) (Wedaman et al., 1996); thus, we chose to target GiKIN2b for morpholino knockdown to compare the resulting phenotype with the known phenotype of the GiKIN2a dominant-negative. To track the levels of the protein, we produced a GiKIN2b-specific antibody that recognizes a single 72-kDa band in Giardia extracts (Figure 2.6A). This antibody was used on immunobots along with an anti-protein-disulfide isomerase 2 (PDI-2) antibody as a loading control (Lauwaet et al., 2007) (Figure 2.6B), and the ratio of these two proteins was used to compare amounts of GiKIN2b at different time points (Figure 2.6B and C). When used at 100 µM, the anti-GiKIN2b morpholino achieved a 60% reduction in protein levels in 24 hours (Figure 2.6C). Mutants with disrupted cytoskeletons were also observed in fixed cells (Figure 2.6D). The cytoskeleton of Giardia cells includes four pairs of flagella, a bundle of microtubules called the median body, and several other structures (Elmendorf et al., 2003)(Figure 2.6D). In the mutant cells, the entire external region of at least two pairs of flagella was missing, with only internal axonemes remaining. The caudal and posteriolateral flagella were most often affected, though three or even all four sets of flagella were affected in some cells. These cells also often had reduced, disorganized, or missing median bodies, suggesting that GiKIN2b may also be involved in regulating microtubule dynamics in these unique giardial structures. However, other microtubule arrays, including the relatively stable ventral disk, were not perturbed. After 24 hours, approximately 14% of cells in the knockdown cultures displayed mutant phenotypes, compared to 5% in the mispair control and 1% in the no morpholino control (Figure 2.6E). The mutant cells seen in the mispair control could reflect the slight ability of this morpholino to knock down GiKIN2b; however, the population-wide kinesin-2 protein levels in these cultures are approximately the same as the no-morpholino control (Figure 2.6B and C).

Using an epitope-tagged protein to measure morpholino knockdown Because translation-blocking morpholinos are the only option for use in Giardia (unless the desired target is one of the four intron-containing genes), determining the level of knockdown generally requires a specific antibody. However, zebrafish researchers have had success using an epitope-tagged protein as a readout for morpholino efficacy (Collart et al., 2005). To determine whether this strategy would work in Giardia, we used the GiKIN2b morpholino in a strain carrying a plasmid containing a C-terminally GFP-tagged GiKIN2b under its native promoter (Hoeng et al., 2008). Thus, the same morpholino could be used to knock down the endogenous and GFP-tagged GiKIN2b simultaneously.

25 Although the endogenous protein in this strain was reduced to the same level as in the wild-type after 24 hours, GiKIN2b::GFP took a total of 48 hours after electroporation to reach that level (Figure 2.7A). This delay is likely due to a difference in turnover rate between the endogenous GiKIN2b and the GFP-tagged protein, and probably also high gene dosage in cells with multiple copies of the plasmid. Twenty-four hours after electroporation, 50% of these cells displayed mutant phenotypes (shortened/missing flagella), (Figure 2.7B). In fact, even the GiKIN2b::GFP strain without morpholino treatment contained 15% mutant cells. We hypothesize that the GiKIN2b::GFP fusion protein interferes with wild-type GiKIN2b complex function; consequently, these cells are sensitized to morpholino knockdown of the remaining GiKIN2b, which results in effectively lower levels of functional complex. In any case, the GFP-tagged protein was eventually knocked down to the same extent as the endogenous protein.

Discussion General considerations for morpholino use in Giardia In this study, we sought to determine whether morpholinos could effectively reduce gene expression by translational inhibition in Giardia intestinalis. We chose to target two genes, GFP and GiKIN2b, so that we could determine the effect of morpholino knockdown of a neutral gene (GFP), and then compare the phenotype of targeting an endogenous gene with that of a previously characterized homolog. Using morpholinos at a final concentration of 100 µM, we achieved a 60% reduction in protein levels in 24 hours for both of these targets. However, a recent study on actin function in Giardia achieved up to 80% knockdown with this method (Paredez et al., 2011), further highlighting the utility of morpholinos on a variety of targets. Differences in the level of knockdown achieved by different morpholinos could be related to the transcript abundance of the target gene, protein turnover, and/or the properties of the individual morpholinos. Although we found that maximum reduction of protein levels was achieved at 100 µM morpholino for the abundantly expressed GFP construct, targets expressed at a lower level could require less morpholino to reach the same level of knockdown. In addition, morpholinos can be designed to target the 5’ untranslated region (UTR) of a gene. While these are, on average, as effective as those targeting the first 25 bases of the coding sequence, the exact location of morpholino binding on the mRNA transcript can alter its efficacy on that specific target (Summerton, 1999). Because Giardia transcripts generally have very short 5’UTRs (0-14 nucleotides) (Li and Wang, 2004), they are difficult to predict and must be determined experimentally, making targeting the coding region a more straightforward option. However, in some cases, the sequence properties of a morpholino targeted to the coding sequence may be undesirable (extensive secondary structure, for example), requiring that a portion of the 5’UTR be targeted instead (Moulton and Yan, 2008). One potential drawback to the use of translation-blocking morpholinos is that the reduction of gene expression must be quantified on the protein level. However, we have shown here that an epitope-tagged version of the protein, if expressed 26 under its native promoter, can be used as a readout for the efficacy of the morpholino on the endogenous protein. Of course, the epitope tag must be placed on the C terminus of the protein of interest, as the translation start site must remain the same as the endogenous protein. In the case of GiKIN2b::GFP strain, the GFP- tagged protein took an additional 24 hours to reach the same level of knockdown as the endogenous GiKIN2b, probably due to a difference in turnover rate caused by the GFP tag. Nevertheless, the levels of the endogenous protein were already reduced by 60% after 24 hours, and cells with the mutant phenotype were abundant at this time. In fact, for this particular protein, the addition of the GFP tag alone actually caused a slight mutant phenotype in the controls not treated with morpholino. The phenotype of these mutants, which had shortened external flagella, is consistent with the phenotype seen after morpholino treatment, suggesting that the GFP- tagged protein can interfere with the normal function of the complex. As a consequence, these cells were hypersensitive to the addition of the morpholino, which dramatically increased the abundance of cells displaying the mutant phenotype. While other epitope-tagged proteins may function more normally, they can still serve as a reporter for the level of knockdown achieved by the morpholino. Furthermore, although we used genes expressed from episomes in the present study, an endogenously tagged gene (Gourguechon and Cande, 2011) could serve as an even more sensitive reporter because it is likely to be expressed at levels closer to that of the endogenous gene.

Insights into kinesin-2 function in Giardia The phenotype resulting from targeting GiKIN2b with morpholinos resembled that of the dominant-negative GiKIN2a (Hoeng et al., 2008), but revealed some aspects of kinesin-2 function that the dominant negative may have obscured. Rigor mutants of kinesins act as dominant negatives due to their ability to bind to the microtubule but not hydrolyze ATP, thereby effectively creating a “roadblock” that also prevents the endogenous kinesin from functioning (Crevel et al., 2004). In the previous study of GiKIN2a, the induction of the dominant-negative GiKIN2a by a tetracycline-repressible vector resulted in a 15-30% decrease in flagellar length in the overall population of transformants (Hoeng et al., 2008). The authors speculated that, in the absence of functional GiKIN2a, the remaining regions of the external flagella may have been maintained by the inclusion of homodimers of GiKIN2b in the complex, as has been seen for kinesin-2 homologs in Tetrahymena (Brown et al., 1999). However, morpholino knockdown of GiKIN2b produced cells missing the entire external regions of their flagella, suggesting that the remaining external flagellar regions in the dominant-negative cells were probably due to residual activity of the wild-type heterotrimeric complex. The disruption of the median bodies, enigmatic arrays of microtubules mainly associated with the cytoplasmic axonemes of the caudal and ventral flagella in interphase Giardia cells, in the morpholino-treated cells was also not seen in the dominant-negative GiKIN2a strain (Hoeng et al., 2008). GFP-tagged GiKIN2a and

27 GiKIN2b both localize primarily to the flagellar tips, the exit points of the flagella from the cell body, and the basal bodies (Brown et al., 1999). Thus, it is unclear whether these proteins have a direct role in median body maintenance or whether the phenotype is an indirect effect, perhaps due to disruption of the flagella. Interestingly, expression of a dominant-negative mutant of kinesin-13, which regulates microtubule dynamics in Giardia and localizes to the flagellar tips and the median body, causes elongation of both the flagella and the median body (Dawson, 2007). Another possibility is that the knockdown of GiKIN2b causes a mitotic delay or arrest—the median bodies disappear during mitosis in Giardia (Sagolla et al., 2006), and kinesin-2 homologs have been shown to play a role in mitosis in other species (Haraguchi et al., 2006; Miller et al., 2005). In any case, some conclusions drawn from both studies are in concordance: that different pairs of flagella are not equally susceptible to kinesin-2 disruption, and that the cytoplasmic regions of the flagella are not maintained solely by kinesin-2 mediated intraflagellar transport, as these were still at least partially present in the mutant cells.

Conclusions Existing technologies for reducing gene expression in Giardia require a period of selection that may cull cells displaying the most extreme phenotypes, thus precluding the disruption of essential genes (Prucca et al., 2008). However, because the time from electroporation to phenotype is only 24 hours on average, morpholinos allow for the knockdown and rapid characterization of essential and non-essential genes alike. Even if knockdown of a certain gene causes rapid cell death, mutants can be recovered by monitoring the growth of cells after electroporation and collecting samples as cells begin to die. Fluorescent modifications can also be added to morpholinos, allowing for the identification of cells containing the highest quantities of morpholino to help uncover more subtle phenotypes. Finally, because morpholinos are stable for several days in cell culture, it may be possible to electroporate cells with morpholinos targeting a gene of interest and then use these cells to infect an animal host, testing the gene’s effect on pathogenicity. Thus, we believe morpholinos have the potential to become a powerful new tool in the field of Giardia biology.

Materials and Methods Culture conditions Giardia intestinalis trophozoites, WB clone C6 (ATCC 50803), were grown in modified TYI-S-33 medium with bovine bile (Sigma-Aldrich, St. Louis, MO)(Keister, 1983). Cultures were incubated in 16 ml plastic screw top tubes (Fisher Scientific, Pittsburgh, PA) at 37°C and passaged every three days.

Strain construction

28 For the GFP construct, the pMCS-eGFP vector was altered to include the promoter of the glutamate dehydrogenase (GDH) gene fused upstream of the eGFP open reading frame (Sagolla et al., 2006; Yee et al., 2000). The GiKIN2b::eGFP strain was a gift from S. C. Dawson (Hoeng et al., 2008). Transformation of Giardia trophozoites was done as previously described (Singer et al., 1998) with modified electroporation conditions for the GenePulserXL (Bio-Rad, Hercules, CA) at 375mV, 1000 µF and 700 ohms. After electroporation, the cells were added to 12 mL of fresh warm growth medium, allowed to recover at 37°C overnight, and then selected with 10 µg/mL puromycin (Calbiochem, La Jolla, CA) until confluent. Strains were maintained at a final concentration of 50 µg/mL puromycin.

Electroporation of morpholinos Electroporation of morpholinos was done essentially as previously described for plasmids (Singer et al., 1998). Briefly, cells ~1 day past confluency (~1–2 x 106 cells/mL) were prepared for electroporation by washing each tube once with 1.5 mL of cold fresh media, and then resuspending in 1.2 mL of cold fresh media. From this solution (~1.6 x 107 cells/mL), 0.3 mL was aliquotted into each 0.4 cm cuvette and kept on ice. Lyophilized morpholinos (Gene Tools, LLC, Philomath, OR) were resuspended in sterile water to a concentration of 1 mM. This stock was added directly to the cuvette with cells to produce the desired concentration of morpholino. For the negative control, the same volume of sterile water was added. The cuvettes were incubated on ice for 15 minutes, electroporated with the conditions described above, and then incubated on ice another 15 minutes. Finally, each cuvette was used to inoculate three 8-mL tubes containing fresh, warm media (containing puromycin for the GFP experiments) by aliquotting 100 µL of electroporated cells into each tube.

Flow cytometry For flow cytometric (FACS) quantitation of eGFP levels, 1 mL of cells from each time point was collected by centrifugation, then resuspended in warm HEPES- buffered saline (HBS) for 30 minutes at 37°C to facilitate eGFP fluorescence. Cells were then fixed with 1% paraformaldehyde for 30 minutes at 37°C, washed twice with HBS, and finally resuspended in 1 mL HBS. Samples in HBS were counted on a Beckman-Coulter EPICS XL Flow Cytometer. 20,000 cells from each sample were counted and eGFP fluorescence measured, and a gate for eGFP positive cells was created based on comparison of the starting eGFP-expressing culture to wild-type cells.

Production of anti-kinesin-2b antibody To produce a polyclonal antibody recognizing GiKIN2b (GenBank accession XP_001708236), a 732-nucleotide segment of the gene containing the C-terminal stalk and tail of the protein was cloned from Giardia genomic DNA using primers kin-2b forward (5’-ACTGATATCTAATGGGTGCAGGGTTTACGGGCTATAC-3’),

29 and kin-2b reverse (5’-ACTGCGGCCGCTCAACCGAAACCAGCCATGCCACG-3’) into the EcoRV and Not1 sites of the vector pET28c (EMD Biosciences, Gibbstown, NJ). This vector contains an N-terminal His-tag to allow for protein purification. The vector was transformed into E. coli BL21(DE3) (EMD Biosciences), and the correct insert in transformants was confirmed by sequencing. Binding to Ni-NTA beads under denaturing conditions was performed according to the manufacturer’s instructions (Qiagen, Valencia, CA). The purified protein was used to inoculate two New Zealand White rabbits according to a 77-day immunization protocol (Covance, Denver, PA). Sera from the final bleed of one rabbit recognized a single band of the appropriate weight (~72 kDa) on immunoblots of Giardia extracts and was used for subsequent analyses.

Immunoblotting and protein quantification Cells from each time point were placed on ice for 15 minutes to detach, collected by centrifugation, and then washed once in cold HBS containing 1X Halt Protease Inhibitor Cocktail (Thermo Fisher Scientific, Rockford, IL) and 5 mM EDTA. The pellet was resuspended in 1x Laemmli buffer minus dithiothreitol and bromophenol blue, boiled for 5 minutes, and then frozen. The protein concentration of each sample was determined using the DC Protein Assay (Bio-Rad, Hercules, CA) according to the manufacturer’s instructions, and all samples were diluted to 2 mg/mL with 2x Laemmli buffer. 10-20 µg of cell extract was loaded onto 10% SDS- PAGE gels and separated by electrophoresis, then transferred to Immobilon PVDF- FL membranes (Millipore, Billerica, MA). Membranes were blocked with a 5% milk/0.5% Tween-20 solution, then probed with one or more of the following antibodies: the rabbit polyclonal anti-GiKIN2b antibody described above at 1:50,000, and a rabbit polyclonal anti-Giardia protein-disulfide isomerase 2 (PDI-2) antibody at 1:500,000, a gift from F. D. Gillin (Knodler et al., 1999). After washing, membranes were then probed with the IRDye 800CW goat anti-rabbit secondary antibody at 1:15,000 (Li-Cor Biosciences, Lincoln, NE). The blots were visualized with a Li-Cor Odyssey Infrared Imager. Densitometry of bands was performed using the Li-Cor Odyssey software according to the manufacturer’s instructions.

Visualization of GFP and the cytoskeleton Direct fixation of GFP and antibody immunostaining (TAT1 antibody (Woods et al., 1989), a gift from K. Gull, to visualize microtubules) were performed as described (Poxleitner et al., 2008). (D'Souza and Michiels, 2010)

Fluorescence deconvolution microscopy Images were collected with SoftWorX image acquisition software (Applied Precision, Issaquah, WA) on an Olympus IX70 wide-field inverted fluorescence microscope with an Olympus PlanApo 60, NA 1.4, oil-immersion objective and Photometrics CCD CH350 camera (Roper Scientific, Tuscon, AZ). Image stacks were deconvolved

30 with the SoftWorX deconvolution software and flattened as maximum projections (Applied Precision, Issaquah, WA).

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Yee, J., Mowatt, M.R., Dennis, P.P., and Nash, T.E. (2000). Transcriptional analysis of the glutamate dehydrogenase gene in the primitive eukaryote, Giardia lamblia. Identification of a primordial gene promoter. J Biol Chem 275, 11432-11439.

33 Table 2.1: Sequences of the morpholinos used in this study. Sequences are shown from 5’ to 3’. Altered nucleotides in the mispair morpholino are shown in lowercase letters. Anti-GFP CAGCTCCTCGCCCTTGCTCACCATG Anti-GFP mispair CAcCTgCTCGCCgTTGCTgACgATG Anti-Kinesin-2b GCCTTTGCCCTTACTCTTGCTCATC Anti-Kinesin-2b mispair GCgTTTcCCCTTAgTCTTcCTgATC

34 Figure 2.1: Time course of GFP knockdown by morpholinos. Cells were collected at the indicated times after electroporation with water (no morpholino control), 100 µM mispair anti-GFP morpholino, 10 µM anti-GFP morpholino, or 100 µM anti-GFP morpholino. Fixed samples were subjected to flow cytometry and categorized as GFP positive or GFP negative compared to a wild-type control (see Figure 2.4). The number of GFP positive cells in the no morpholino control from each time point was set to 100%. The 96 hour culture consisted of cells that were passaged 72 hours after electroporation, then grown an additional 24 hours before collection. These data are the average of three biological replicates, and error bars represent one standard deviation.

35 36 Figure 2.2: Illustration of gating parameters used in Figure 2.1. Representative flow cytometry histograms demonstrating the gating parameters used to distinguish “GFP positive” cells from “GFP negative” cells in Figure 2.1. These histograms are from the 24-hour time point. The gate for GFP fluorescence was drawn based on the WT negative control (upper left). The percentages of cells in each gate are indicated in the upper left and upper right corners of the histogram.

37 WT 10 uM anti-GFP morpholino

Water control 100 uM anti-GFP morpholino

Mispair morpholino

38 Figure 2.3: Representative histograms of flow cytometry data presented in Figure 2.1. The experiment was performed as described in Figure 2.1. The gates shown in Figure 2.1 were used to determine the percentages of GFP positive and GFP negative cells enumerated in Figure 2.1. The amount of GFP fluorescence in the negative controls [water (no morpholino) control and mispair morpholino control] decreases somewhat over time (48h and 72h time points); this is likely due to negative regulation of the glutamate dehydrogenase promoter as the cells reach confluency and the stationary phase of growth.

39 40 Figure 2.4: Doubling the concentration of morpholinos does not result in a substantial increase in levels of knockdown. (A) Representative flow cytometry histograms of cells that were collected at the indicated times after electroporation with 200 µM mispair anti-GFP morpholino, 100 µM anti-GFP morpholino, or 200 µM anti-GFP morpholino. Fixed samples were subjected to flow cytometry and categorized as GFP positive or GFP negative as described in Figure 2.3. (B) A graphical representation of data from two experiments like the one in (A). The number of GFP positive cells in the mispair control from each time point was set to 100%.

41 A

B

42 Figure 2.5: Efficiency of electroporation of morpholinos at different concentrations. Twenty-four hours after electroporation, >99% of cells are positive for fluorescent mopholinos at 100 µM and 200 µM concentration, but only ~37% are positive at 10 µM morpholino. GFP-expressing cells were electroporated as described in the text, but using an anti-GFP morpholino with a red fluorescent end modification (lissamine) or a non-fluorescent mispair control. Flow sorting was performed on live cells, as fixation of the fluorescent morpholino is not recommended by the manufacturer. Cells are plotted with GFP fluorescence on the x-axis and morpholino fluorescence on the y-axis. Gates were set based on wild-type cells (upper left plot). The horizontal line is the gate between “morpholino positive” and “morpholino negative” cells, while the vertical line separates “GFP positive” and “GFP negative” cells. The percentage of cells in each quadrant is indicated in the four corners of the plots. All five plots are combined in the bottom right graph. We did not find any obvious negative correlation between levels of morpholino fluorescence and GFP fluorescence within a given morpholino-treated population, but GFP knockdown does increase in a concentration-dependent manner.

43 WT 100 uM anti-GFP morpholino

Mispair control 200 uM anti-GFP morpholino

10 uM anti-GFP morpholino

44 Figure 2.6: Translation-blocking morpholinos reduce GiKIN2b protein levels and cause extreme shortening of the flagella. (A) The GiKIN2b antibody produced for this study recognizes a band of the correct size (72 kDa) on Giardia extract immunoblots. (B) Results from a representative immunoblot of samples 24 hours after electroporation with water (no morpholino control), 100 µM mispair anti-GiKIN2b morpholino, or 100 µM anti-GiKIN2b morpholino. Protein-disulfide isomerase 2 (PDI-2) was used as a loading control. For each sample, the ratio of GiKIN2b to PDI-2 is given. (C) An average of the ratio of GiKIN2b to PDI-2 on immunoblots from three experiments like the one in (B). The GiKIN2b:PDI-2 ratio in the no morpholino control was set to 100%. Error bars represent one standard deviation from the average of three biological replicates. (D) A diagram of a Giardia cell, and immunolabeling of wild-type and anti-GiKIN2b morpholino-treated cells. Fixed cells were labeled with an anti-α-tubulin antibody. Scale bar = 11 µm. In the diagram, the identities of the four pairs of flagella are indicated: afl, anterior flagella; pfl, posteriolateral flagella; vfl, ventral flagella; cfl, caudal flagella. The median body (mb, a bundle of microtubules with unknown function), ventral disk (vd, used to attach to substrates), and two nuclei (N) are also labeled. (E) The distribution of mutant phenotypes in fixed and immunolabeled samples. Cells were collected 24 hours after electroporation with water (no morpholino control), 100 µM mispair anti-GiKIN2b morpholino, or 100 µM anti- GiKIN2b morpholino. Cells were classified as normal, mutant (missing the external portion of at least two pairs of flagella), or other (cannot be categorized). 100 cells were counted for each sample.

45 46 Figure 2.7: GFP-tagged GiKIN2b is knocked down by the anti-GiKIN2b morpholino. (A) The ratio of GiKIN2b or GiKIN2b::GFP to PDI-2 from immunoblots is plotted, with the mispair control set to 100%. Cells were collected 24 and 48 hours after electroporation with 100 µM mispair anti-GiKIN2b morpholino or 100 µM anti- GiKIN2b morpholino. For each time point, the first bar is the endogenous GiKIN2b in the mispair-treated cells, the second bar is the endogenous GiKIN2b in the morpholino-treated GiKIN2b::GFP strain, and the third bar is the GFP-tagged GiKIN2b, which is also recognized by the GiKIN2b antibody, in that strain. (B) The distribution of mutant phenotypes in fixed and immunolabeled samples. GiKIN2b::GFP cells were collected 24 hours after electroporation with water (control) or 100 µM anti-GiKIN2b morpholino. Cells were classified as normal, mutant (missing the external portion of at least two pairs of flagella), or other (cannot be categorized). 100 cells were counted for each sample.

47 48 Chapter 3

Nuclear behavior and the cytoskeleton during differentiation of Giardia intestinalis

Abstract The protozoan intestinal parasite Giardia intestinalis is a major waterborne pathogen. During its life cycle, Giardia alternates between the actively growing trophozoite stage, which has two diploid nuclei with very low levels of allelic heterozygosity, and the infectious cyst stage, which has four nuclei and a tough outer wall. Although the transition to the cyst has been studied extensively with respect to wall formation, we still lack basic knowledge about the changes that occur inside the cyst, including the origin of the four nuclei. In this study, we used immunofluorescence staining to track the major components of the microtubule cytoskeleton, including the flagella, basal bodies, and adhesive disk, during encystation and excystation. We are the first to directly demonstrate that the Giardia cyst is formed from a single trophozoite by a mitotic division without cytokinesis, as opposed to the fusion of two trophozoites. The cell pauses after completing mitosis, with four nuclei and two partially formed adhesive disks. During excystation, division resumes, and the cell completes cytokinesis to form two daughter trophozoites. In addition, we show that the original nuclear pairs from the parent trophozoite remain associated in the cyst and are distributed as pairs to daughter cells during excystation. Thus, targeted nuclear sorting (such that each daughter cell receives a pair of identical nuclei) does not appear to be a method by which Giardia reduces heterozygosity between nuclei. These results shed light on basic aspects of the Giardia life cycle and have implications for our understanding of the population genetics of this parasite.

Introduction The divergent eukaryotic parasite Giardia intestinalis (syn. Giardia lamblia, Giardia duodenalis) is a major cause of diarrheal disease worldwide. The Giardia life cycle consists of two stages: a flagellated trophozoite with two nuclei and an infectious cyst with four nuclei. The two diploid nuclei in the trophozoite appear to be genetically identical and are both transcriptionally active (Bernander et al., 2001; Kabnick and Peattie, 1990b; Yu et al., 2002). In addition, they remain independent in the trophozoite, dividing with separate spindles by a semi-open mitosis, and one copy of each parental nucleus is inherited by each daughter cell (Ghosh et al., 2001; Sagolla et al., 2006; Yu et al., 2002). The differentiation from trophozoite to cyst occurs when a trophozoite, which is usually found attached to the wall of the small intestine, is swept towards the large intestine. Changes in pH and cholesterol availability prompt the trophozoite to encyst, forming a cyst with four nuclei and a thick outer wall (Gillin et al., 1989; Lujan et al., 1996; Lujan et al., 1997). The formation of the cyst wall, which involves the regulated secretion of cyst wall protein components, has been a topic of intense 49 study over the past few decades (see (Lauwaet et al., 2007) and references therein). The cyst is water resistant and can persist for weeks in the environment until it is ingested by a new host. After passing through the stomach, the cyst undergoes excystation, releasing an “excyzoite” that rapidly divides to produce two daughter trophozoites (Bernander et al., 2001; Buchel et al., 1987). In vitro, encystation can be prompted by exposing cells to basic media containing high levels of bile (which prevents cholesterol absorption) for 24-48 hours (Gillin et al., 1987; Gillin et al., 1989; Gillin et al., 1988; Kane et al., 1991; Lujan et al., 1996). Similarly, excystation can be triggered by two-stage procedure in which cells are first exposed to acidic conditions for 30 minutes, mimicking passage through the stomach, followed by a higher-pH solution containing proteases for an addition 30 minutes, which helps break open the cyst wall (Bingham and Meyer, 1979; Boucher and Gillin, 1990; Buchel et al., 1987; Hautus et al., 1988; Rice and Schaefer, 1981; Schaefer et al., 1984). While many advances have been made in our understanding of cyst wall formation, the cytoskeletal changes underlying both encystation and excystation have received less attention, with studies focusing primarily on ultrastructural aspects of the transition and on cyst wall morphology (Bittencourt-Silvestre et al., 2010; Buchel et al., 1987; Chavez-Munguia et al., 2007; Erlandsen et al., 1989; Hetsko et al., 1998; Lanfredi-Rangel et al., 2003; Luchtel et al., 1980; Midlej and Benchimol, 2009; Palm et al., 2005; Sheffield and Bjorvat, 1977). The trophozoite microtubule cytoskeleton consists of four pairs of flagella, with eight basal bodies located between the two nuclei, as well as a ventral adhesive disk (used to attach to the intestinal epithelium) and a prominent bundle of microtubules of unknown function called the median body (Elmendorf et al., 2003). These structures are largely disassembled during cyst formation, with only internal axonemes and disk fragments remaining in the mature cyst (Elmendorf et al., 2003). It has been assumed that the cyst is formed via an incomplete mitotic division (as opposed to the fusion of two trophozoites); however, to our knowledge, this has never been directly demonstrated (Fig. 1), although studies have suggested that an encystation restriction point exists at the G2 phase of the cell cycle (Reiner et al., 2008). Related to the question of how the quadrinucleate cyst is formed are the identities of the nuclei in the cyst and during excystation. The Giardia genome contains very low levels of heterozygosity (Franzen et al., 2009; Jerlstrom-Hultqvist et al., 2010; Lasek-Nesselquist et al., 2009; Morrison et al., 2007; Teodorovic et al., 2007), suggesting that the two genomes are able to exchange genetic information during the Giardia life cycle. However, another way to maintain heterozygosity could be the sorting of nuclei during excystation such that each daughter cell receives two identical nuclei (rather than one of each original nucleus from the parent trophozoite)(Figure 3.1). Thus, determining how the nuclei are inherited during differentiation has implications for our understanding of both the basic biology and the population genetics of this parasite. In the present study, we used immunofluorescence staining of the cytoskeleton and cyst wall to determine how the

50 cyst is formed and to characterize the pattern of nuclear inheritance during encystation and excystation.

Results Cysts are formed by an incomplete mitotic division To determine whether the cyst is derived from a mitotic division or fusion of two trophozoites, we first mixed two strains with different epitope-tagged markers (histone 2A-3HA and median body protein-3myc) and encysted them in vitro. None of the resulting cysts contained both markers (Table 3.1). Following water treatment (to destroy any unencysted cells), excystation, and growth to confluency of these mixed cultures, we found that trophozoites always had only one of the two markers (Table 3.1). This result indicates that, at least in vitro, cysts are derived from a single trophozoite. Additionally, in a larger-scale experiment performed with 5 x 107 live cysts (500-mL cultures), encystation and excystation of a mixture of two strains with different integrated drug resistance markers (neomycin and puromycin) failed to produce any trophozoites with resistance to both drugs (data not shown). Because we typically observe an in vitro excystation efficiency on the order of ~1%, the production of recombinant cells (which would require cell fusion followed by chromosomal recombination or nuclear sorting), if it occurs, would have to happen in fewer than 1 in 500,000 cells. Thus, we sought to characterize the mitotic division during encystation by staining cells with tubulin, centrin, and cyst wall protein (CWP) antibodies after 24 hours of encystation (Figure 3.2). Prior to the division, the trophozoites are observed producing encystation-specific vesicles (ESVs) full of CWP, and two pairs of four centrin spots (marking the basal bodies) are visible between the nuclei (Figure 3.2A). Anaphase (Figure 3.2B) and telophase (Figure 3.2C) generally resemble the mitotic division in trophozoites, with two parallel spindles and the basal bodies/centrioles at the spindle poles. However, unlike in the trophozoite, the encysting cells are concurrently in the process of exporting ESVs to the periphery of the cell for deposition. At the end of telophase of the encystation mitotic division (Fig. 2D), the process diverges further from its vegetative counterpart: cytokinesis does not occur, the cell does not start to grow new flagella, and it begins to disassemble and/or resorb its existing flagella. In addition, the parental adhesive disk is disassembled and two new discs begin to be made (as they would be during a vegetative division (Tumova et al., 2007)) (Figure 3.2E), but these are arrested before growth is complete (see below). At this stage, the two pairs of nuclei are found at opposite poles of the cell (approximately at the previous location of the spindle poles) and remain associated with several bundles of internal axonemes, which appear to be derived from the axonemes that remain between the nuclei during mitosis (Nohynkova et al., 2006) (Figure 3.2E). It is at this time that the cyst wall is rapidly deposited—by the time the pairs of nuclei have migrated to the same pole of the cyst, the cyst wall has been completely formed. The nuclear pairs are closely associated and remain motile for some time in the cyst (Poxleitner et al., 2008), possibly due to their association with 51 the internal axonemes, and this is presumably the mechanism by which they migrate together. The mature cyst contains four nuclei, two bundles of internal axonemes with eight centrin loci, and several ventral disk fragments (Figure 3.2F).

Generation of disc fragments in the cyst It has been reported that the mature cyst contains four ventral disk fragments derived from the disassembly of the parental disk, and it was suggested that these are reassembled during excystation (Elmendorf et al., 2003; Palm et al., 2005). To further investigate the fate of the disk during encystation and excystation, we stained encysting and excysting cells with antibodies to tubulin (Woods et al., 1989) and delta-giardin (Jenkins et al., 2009), a component of the disk (Figure 3.3). Using tubulin staining, we observed the initial stages of two new daughter discs being formed during the later stages of the encystation mitotic division (Figure 3.3A). At the same time, the parental disk is being disassembled, as has been described in the trophozoite (Tumova et al., 2007). We most often saw two disk fragments in the mature cyst (Figure 3.3B); however, in some cases, up to four fragments were visible, although they were still usually found in two clusters (Figure 3.3C). Therefore, the disk fragments may not represent two whole pieces, but rather bundles of multiple disk components. Interestingly, these fragments were not always located near the nuclear pairs and basal bodies, despite the close association between the basal bodies and disk in the trophozoite. Nevertheless, after excystation, these disk fragments again became associated with the nuclear pairs (and, by extension, the basal bodies) and appeared to be in the correct position and orientation to complete their development into two new discs (Figure 3.3D). As demonstrated below, both daughter discs are completely formed before the excyzoite completes cytokinesis.

Reconstruction of the cytoskeleton during excystation Next, we stained the cytoskeleton of excysting cells to monitor the transition from excyzoite to trophozoite (Figure 3.4). As previously reported, the excyzoite emerges from a single pole of the cyst, which is weakened by the exposure to acidic conditions in the stomach and intestinal proteases (Boucher and Gillin, 1990; Buchel et al., 1987; Coggins and Schaefer, 1986; Hetsko et al., 1998) (Figure 3.4A). The flagella emerge first, followed by the body of the cell (Figure 3.4B). At this stage, the cytoskeleton of the excyzoite still resembles that of the cyst, with internal bundles of axonemes, two pairs of nuclei, and eight centrin spots (Figure 3.4A-C). These cells sometimes display one or several structures that resemble ESVs (arrow in Figure 3.4C); we hypothesize that these result from the residual translation of cyst wall protein transcripts, which are extremely abundant during encystation. The excyzoite immediately begins to duplicate and/or cluster its axonemes, producing two prominent bundles, each associated with a nuclear pair (Figure 3.4C). Although this configuration of axonemes is different from that of a trophozoite, the cell is still capable of swimming (Buchel et al., 1987; Hautus et al., 1988). After additional rearrangements, the excyzoite begins to resemble a heart- 52 shaped (pre-cytokinesis) cell that is essentially indistinguishable from that of a dividing trophozoite, with the exception that each nucleus contains a 4N (tetraploid) DNA content (Figure 3.4E-F). By this stage, the number of basal bodies has increased from 8 to 16, and the cell and rapidly completes the construction of two ventral discs (Figure 3.4E-F). Following cytokinesis, the resulting 8N trophozoites presumably divide without an intervening S phase to produce two 4N trophozoites (Bernander et al., 2001).

Discussion As an organism with two nuclei, Giardia faces unique challenges in coordinating nuclear inheritance throughout its life cycle. Although the functional equivalence of the nuclei in Giardia and their independent distribution during mitosis in the trophozoite are relatively well established (Kabnick and Peattie, 1990a; Yu et al., 2002), their fates during encystation and excystation have only been studied with respect to genome ploidy (Bernander et al., 2001), despite the significance of this knowledge to our understanding of patterns of allelic heterozygosity and genetic diversity in Giardia populations. Thus, the goal of the present study was to determine how the four nuclei are derived in the cyst and how these nuclei are distributed during excystation. Based on strain mixing experiments (Table 3.1) and immunofluorescence staining of encysting cells (Figure 3.2), we have demonstrated for the first time that the quadrinucleate cyst is produced via an incomplete mitotic division that occurs rapidly during the final stages of cyst wall deposition. Additionally, we have provided the first glimpse into the temporal organization of the microtubule cytoskeleton during encystation and excystation. While the fusion of two trophozoites to form a cyst is still theoretically possible, this event would have to be rare and/or happen only under certain conditions (such as in the host). However, given that encystation takes place during transit through the small and large intestine, we speculate that it would be rare for two cells to find each other, and a requirement for cell fusion for cyst formation would be consequently be disadvantageous; thus, encystation from a single parent cell appears to be the predominant (if not sole) mode of cyst formation. Our observations of the cytoskeletal components of encysting and excysting cells both synthesize and build upon the results of previous ultrastructural and TEM studies (Bernander et al., 2001; Bittencourt-Silvestre et al., 2010; Buchel et al., 1987; Hetsko et al., 1998; Luchtel et al., 1980; Midlej and Benchimol, 2009; Palm et al., 2005; Sheffield and Bjorvat, 1977), which have primarily focused on mature cysts. We found that the encystation division resembles the vegetative trophozoite mitotic division until approximately telophase. However, instead of continuing on into cytokinesis, the cell retracts/depolymerizes its flagella and the cyst wall rapidly forms. At the same time, cyst wall components are being produced and transported to the periphery of the cell, a process that appears to be independent from, albeit simultaneous with, the nuclear division (Stefanic et al., 2009). In addition, although trophozoites in the early stages of encystation display

53 larger, rounded ESVs, by the time the nuclear division takes place, the cyst wall components have been transported to the periphery of the cell and exist in a more diffuse network (Konrad et al., 2010). Based on the pattern of centrin and tubulin staining in our study, the bundles of axonemes associated with each nuclear pair in the cyst appear to be derived from the spindle-associated axonemes found associated with the nuclei during mitosis (Nohynkova et al., 2006). These results are consistent with those of a TEM-based study that reported two bundles of three internal axonemes associated with each nuclear pair in the cyst, as well as two additional flagella found in the periphery of the cell (Sheffield and Bjorvat, 1977). Thus, a total of eight basal bodies are present in the mature cyst, which is consistent with our observations. Also found in the mature cyst are two to four fragments of the ventral disk (Figure 3.3). It has been previously suggested that these fragments are derived from the single ventral disk of the parent trophozoite, which is disassembled during encystaton and reassembled during excystation (Palm et al., 2005). However, based on our observations of the growth of two nascent daughter disks during the encystation mitotic division, as well as the overall morphological similarity of these nascent disks to the fragments in the cyst, we hypothesize that disk growth is merely paused during encystation and then resumed during excystation. This scenario would be more consistent with a what is observed during cytokinesis in the trophozoite, in which two new ventral discs grow from the basal bodies on the opposite side of the cell from the parental ventral disk (Tumova et al., 2007). This is also the first study to examine the Giardia cytoskeleton during excystation. As reported in previous studies, we observed that the excyzoite exits the cyst at one pole, starting with its flagella. The excyzoite has a similar cytoskeletal organization to the cyst, but rapidly re-molds its cytoskeleton to resemble a cytokinetic trophozoite, which is the stage where it paused after the mitotic division during encystation. At this stage, each nascent trophozoite has an 8N genetic content (Bernander et al., 2001); thus, after cytokinesis is completed, the cell is primed to complete another round of mitosis without an intervening S phase. Because the number of centrin spots increases from 8 in the excyzoite to 16 in the heart-shaped cells, we conclude that basal body duplication is also occurring during this time. Finally, our observation that nuclear pairing is maintained during encystation and excystation suggests that daughter trophozoites receive one copy of each parental nucleus. To date, the genomes of three Giardia assemblages (distinct genotypes that have been argued to represent different species) have been sequenced, and all have revealed relatively low levels of allelic heterozygosity, ranging from less than 0.01% in assemblages A and E to 0.5% in assemblage B (Franzen et al., 2009; Jerlstrom-Hultqvist et al., 2010; Morrison et al., 2007). These results are surprising because, in a presumed asexual organism with two independent nuclei, the nuclear genomes—and perhaps even homologous chromosomes—would be expected to diverge over time due to drift (Birky, 2010).

54 The distribution of identical nuclei to daughter cells during excystation would explain the low levels of heterozygosity observed in Giardia populations. Thus, our observation that nuclei appear to be distributed in non-identical pairs during excystation is significant. However, while this is the most parsimonious conclusion based on our observations, future studies using integrated markers will be necessary to demonstrate conclusively that the nuclear pairs are not exchanged prior to their distribution during excystation. In conclusion, the maintenance of two equivalent nuclei in Giardia is one of the most intriguing aspects of its biology, and understanding this ability will shed light on both the population genetics and basic biology of this important parasite.

Materials and Methods Culture conditions Giardia intestinalis trophozoites, WB clone C6 (ATCC 50803), were grown in modified TYI-S-33 medium (Keister, 1983) with bovine bile (Sigma-Aldrich, St. Louis, MO). Cultures were incubated in 16-mL plastic screw top tubes (Fisher Scientific, Pittsburgh, PA) at 37°C and passaged every three days.

In vitro encystation Encystation was performed essentially as described (Kane et al., 1991). Briefly, trophozoites were seeded into 30- or 50-mL tissue culture flasks (BD Biosciences, Bedford, MA) and grown to 80%-100% confluency. The media was then decanted and replaced with prewarmed encystation media (10 mg/mL bovine/ovine bile, pH 7.8). After 24 hours at 37°C, cells were either fixed for observation or incubated on ice for 15 min, pelleted at 500 xg, and resuspended in regular growth medium (0.5 mg/mL bile, pH 7.1). After an additional 24 hours of growth at 37°C, the cells were again incubated on ice, pelleted, and resuspended in 50 mL ice-cold, sterile deionized water to destroy all unencysted cells. Cysts were stored at 4°C for 1-3 days, then excysted as described below.

In vitro excystation Excystation was performed essentially as described (Boucher and Gillin, 1990). Briefly, water-treated cysts were pelleted in a 50-mL conical tube, then resuspended in 10 mL prewarmed stage I solution (1X Hank’s Balanced Salt Solution [Invitrogen, Carlsbad, CA], 57 mM L-cysteine HCl, 32.5 mM reduced glutathione, and 25mM NaHC03; the pH was adjusted to 4.0 with 0.01N HCl). The tube was vortexed briefly, then incubated at 37°C for 30 minutes. After another brief vortexing, cells were pelleted at 500 x g for 5 minutes. The supernatant was then decanted and replaced with 10 mL prewarmed stage II solution (Tyrode’s buffer [Sigma-Aldrich, St. Louis, MO] containing 1 mg/mL Trypsin II-S [Sigma]; the pH was adjusted to 8.0 with 1 M NaHC03). The tube was vortexed briefly, incubated at 37°C for 30 minutes, then vortexed again and pelleted. The supernatant was

55 decanted and replaced with prewarmed growth media (see Culture conditions), and cells were moved to 16-mL culture tubes and incubated at 37°C. Cells were then either fixed 1-3 hours later for immunofluorescence staining or grown to confluency (approximately 5-7 days).

Creation of tagged histone 2A and median body protein (MBP) constructs for integration For the histone 2A-3myc strain, part of Giardia ORF GL50803_14256 was amplified from purified WBC6 genomic DNA using the primers H2A-for (5’- TATACTAGTAGCACCAAGCCCGTCAAGGAC-3’) and H2A-rev (5’- TCTCTTAAGAAGATCCTGCGACCGATGG-3’). The resulting fragment was cut with the enzymes SpeI and AflII and inserted between those same sites in the vector pKS-3myc-NEO (Gourguechon and Cande, 2011). This plasmid was then linearized for integration by cutting within the H2A sequence using XhoI. To create the MBP-3HA strain, Giardia ORF GL50803-16343 was amplified from purified WBC6 genomic DNA using the primers MBP-for (5’ATGTCCGAGGCTATGGTTTTC-3’) and MBP-rev (5’- TACCCGGGGAACTTACCACTTCGGCCCATG’3’). The resulting fragment was cut with the enzymes XbaI and SmaI and inserted between those same sites in the vector pKS-3HA-NEO (Gourguechon and Cande, 2011). This plasmid was then linearized for integration by cutting within the MBP sequence using MfeI. Transformation and selection of Giardia trophozoites was done as previously described (Gourguechon and Cande, 2011).

Immunostaining Fixation and immunostaining were performed essentially as described (Carpenter and Cande, 2009). Briefly, cells were fixed in 1X PEM buffer (100 mM PIPES, 1 mM EGTA, 0.1 mM MgSO4) containing 2% paraformaldehyde. After fixation for 30 minutes at 37°C, cells were washed once with PEM buffer, then resuspended in PEM buffer and attached to poly-L-lysine-coated coverslips for 30 minutes. Alternatively, for delta-giardin antibody staining, cells were fixed for 3 minutes in ice-cold methanol, then washed twice in PBS before being attached to poly-L-lysine coverslips. Fixed cells were permeabilized with 0.1% Triton-X-100 for 15 minutes, washed three times with PEM, and blocked in PEMBALG (PEM plus 1% BSA, 0.1% NaN3, 100 mM lysine and 0.5% gelatin from cold water fish skin [Sigma]) for 2 hours. Primary antibody incubations were conducted at room temperature overnight, and antibodies were diluted in PEMBALG as follows: mouse anti- trypanosome alpha-tubulin antibody TAT1, 1:50 ((Woods et al., 1989), a gift from K. Gull); 20H5 mouse anti-centrin antibody, 1: 100 (Salisbury et al., 1988); rabbit anti- sea urchin tubulin antibody 1:25 (Fujiwara and Pollard, 1978); rat anti-HA antibody 1:100 (Roche, Basel, Switzerland); rabbit anti-c-myc antibody 1:100 (Cell Signaling Technology, Danvers, MA); rabbit anti-delta-giardin antibody 1:100 ((Jenkins et al., 2009), a gift from M. Jenkins). After washing with PEMBALG, cells were incubated with secondary antibodies (goat anti-mouse 555 and goat anti-rabbit 56 488 [Invitrogen]) at room temperature for two hours. For labeling with the anti-cyst wall protein antibody, cells were then washed three times with PEMBALG, fixed for 10 minutes in 1% paraformaldehyde, washed three times with PEM, and blocked for an additional 30 minutes in PEMBALG. The cells were then incubated with the Cy5-labeled mouse anti-cyst wall protein antibody (Waterborne, Inc., New Orleans, LA) for two hours. After a final three washes each in PEM and PEMBALG, coverslips were mounted onto slides with Prolong Gold Antifade Solution with DAPI (Invitrogen).

Fluorescence deconvolution microscopy Images were collected with SoftWorX image acquisition software (Applied Precision, Issaquah, WA) on an Olympus IX70 wide-field inverted fluorescence microscope with an Olympus UPlanApo 100x, NA 1.35, oil-immersion objective and Photometrics CCD CH350 camera cooled to -35°C (Roper Scientific, Tuscon, AZ). Serial sections were acquired at 0.2 µm intervals, and data stacks were deconvolved using the SoftWorX deconvolution software. For presentation purposes, two- dimensional maximum intensity projections were created from the 3D data sets using the DeltaVision image analysis software (Applied Precision).

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61 Figure 3.1: Overview of alternative theories for cyst formation and nuclear distribution during excystation. While it has often been assumed that the Giardia cyst is formed via an incomplete mitotic division (A), the alternative route of cell fusion between two trophozoites (B) has never been excluded. In addition, during excystation, two possible routes for the partitioning of nuclei to daughter cells would have dramatically difference consequences for Giardia populations. If each daughter cell receives one copy of each parental nucleus (C), any nuclear heterozygosity is maintained throughout the life cycle. In contrast, if the nuclei were distributed such that each daughter cell received two identical nuclei (D), heterozygosity would be eliminated.

62 Encystation Excystation

A C

OR mitosis OR

B D

cell fusion

63 Table 3.1: Cysts are not formed by the fusion of two trophozoites. Strains containing different markers (median body protein-3HA or histone 2A- 3myc) in approximately equal numbers were encysted and excysted together. Cysts and excysted trophozoites were scored for the presence of each marker.

Stage Marker 1 Marker 2 Both

(median body protein) (histone 2A) markers

Cysts (n=100) 49% 51% 0%

Excysted trophozoites 59.8% 40.2% 0%

(n=500)

64 Figure 3.2: The Giardia cyst is formed by an incomplete mitotic division. Encysting cells were stained with DAPI to label DNA (blue) and antibodies to label tubulin (green), centrin (red), and cyst wall protein (CWP) (grey). (A) An encysting trophozoite, demonstrating typical centrin and tubulin staining and the presence of two nuclei. The median body, a cluster of microtubules of unknown function, is indicated (arrow). (B) An encysting cell in anaphase. The two spindles (arrowheads) and median body (arrow) are indicated. (C and D) Encysting cells in early and late stages of telophase. (E) Early cyst in which the nuclear pairs are still at opposite ends of the cell. The arrow indicates what appears to be a nascent disk fragment associated with one pair of nuclei. (F) Mature cyst in which the nuclear pairs have migrated to one pole of the cell. Scale bar = 5 µm.

65 Centrin Tubulin Merge w/DAPI CWP A Encystation

B

C

D

E

F

66 Figure 3.3: Development of the adhesive disk during encystation and excystation. Encysting and excysting cells were stained with DAPI to label DNA (blue) and antibodies to label cyst wall protein (red) and either tubulin (A) or delta-giardin (B- D) (green). (A) An encysting cell in telophase showing the growth of a new daughter disk (arrowhead) and the disassembly of the parental disk (arrow). (B) A mature cyst containing two disk fragments. (C) An excysting cell showing that each disk fragment appears to be composed of two smaller pieces. (D) An excyzoite in which the disk fragments are again associated with the nuclear pairs and are in the correct orientation to resume their development into complete discs. Scale bar = 5 µm.

67 A B C D

tubulin δ-giardin δ-giardin δ-giardin Encystation Excystation

68 Figure 3.4: Reorganization of the microtubule cytoskeleton during excystation. Excysting cells were stained with DAPI to label DNA (blue) and antibodies to label tubulin (green), centrin (red), and cyst wall protein (CWP) (grey). (A) Cyst in the early stages of excystation (note the rupture in cyst wall at the bottom of the image). (B) An excyzoite emerging from a cyst. (C) Completely excysted cell. A residual encystation-specific vesicle is indicated (arrow). (D and E) Later-stage excyzoites in which the typical heart-shaped cytokinetic morphology is restored, including two complete discs and 16 flagella. Scale bar = 5 µm.

69

70

E

D

C B

C

A

Excystation

Tubulin Merge Merge Centrin Chapter 4

Nuclear fusion and genetic exchange between nuclei in the Giardia intestinalis cyst

Abstract The diplomonad parasite Giardia intestinalis contains two functionally equivalent nuclei that are inherited independently during mitosis. Although presumed to be asexual, Giardia has low levels of allelic heterozygosity, indicating that the two nuclear genomes may exchange genetic material. The Giardia genome also contains homologs of meiotic genes (HMGs) that in other organisms are required specifically for meiotic homologous recombination. In the present study, we set out to determine when and where the HMGs are expressed during the life cycle and to test for genetic exchange between nuclei. Three of the HMGs (Spo11, Dmc1a, and Hop1) are expressed only during encystation, while Dmc1b is only expressed in the vegetative trophozoite, suggesting that, despite its annotation as a Dmc1 homolog, it may play a Rad51-like function in somatic DNA damage repair. Mnd1 and Hop2 are expressed at low levels in both trophozoites and cysts, indicating that they may also play a more general role in recombination in Giardia. Using strains containing integrated chromosomal markers, we demonstrate that nuclear fusion and genetic exchange occur between nuclei in the cyst, a parasexual process we have termed “diplomixis.” At the same time, the nuclear pairs and their associated bundles of axonemes exhibit dynamic movements, which may be the driving force for the fusion event. Based on their timing of expression, we hypothesize that the HMGs are involved in this genetic exchange, presumably mediating homologous recombination between the nuclei. If accompanied by mitotic recombination and gene conversion, diplomixis could be a mechanism by which Giardia reduces heterozygosity between its two nuclear genomes.

Introduction Giardia intestinalis is a common intestinal protozoan parasite and a major cause of water-borne diarrheal disease worldwide (Savioli et al., 2006). As a diplomonad, it has two apparently genetically identical, functionally equivalent diploid nuclei (Bernander et al., 2001; Kabnick and Peattie, 1990; Yu et al., 2002). Both nuclei are transcriptionally active and contain two complete copies of the genome. (Kabnick and Peattie, 1990; Yu et al., 2002) The parasite has two developmental stages: a binucleate, flagellated trophozoite that attaches to the upper small intestine, and the quadrinucleate infectious cyst, which is excreted from the body and can persist for weeks in the water supply. The two nuclei in the trophozoite remain physically and genetically distinct during mitosis, with two autonomous spindles segregating the parental nuclei into the daughter cells (Sagolla et al., 2006). As demonstrated in Chapter 3, during encystation, the two nuclei undergo a nuclear division without a subsequent cytokinesis, giving rise to a mature cyst with four nuclei. Upon excystation, the cell 71 is thought to go through one cellular division and then one nuclear and cellular division to form four binucleate trophozoites (Bernander et al., 2001). Neither mating nor meiosis has been reported in Giardia, although several studies have found evidence of genetic exchange in wild Giardia populations (Cooper et al., 2007; Lasek-Nesselquist et al., 2009). Interestingly, the Giardia genome contains seven homologs of meiosis-specific genes (HMGs): Spo11, Hop1, Hop2, Mnd1, Dmc1a, Dmc1b, and Mer3 (Malik et al., 2008; Ramesh et al., 2005). The two Dmc1 homologs were annotated based on phylogenetic analysis, and no Rad51 homolog was annotated in these studies (Malik et al., 2008; Ramesh et al., 2005). If Giardia is indeed asexual and its nuclear genomes remain independent throughout the life cycle, it would be expected to accumulate substantial allelic heterozygosity within and between the two nuclei (Birky, 2005). However, several studies have reported very low levels of allelic heterozygosity in the genome, ranging from <0.01% to 0.5% in different genotypes (Franzen et al., 2009; Jerlstrom- Hultqvist et al., 2010; Morrison et al., 2007). In addition to its importance as a parasite, Giardia is also a divergent eukaryote, belonging to the earliest-diverging eukaryotic lineage in many rooted phylogenies (Arisue et al., 2005; Baldauf et al., 2000; Ciccarelli et al., 2006; Morrison et al., 2007). As such, the roles of the HMGs in Giardia could shed light on their functions early in eukaryotic evolution. It is generally accepted that the machinery for homologous recombination during meiosis was co-opted from DNA damage repair machinery (Marcon and Moens, 2005; Villeneuve and Hillers, 2001). Is Giardia using the HMGs for homologous recombination during a sexual (or parasexual) process, or is it using them solely for DNA damage repair? To address this question, we first determined when and where the HMGs are expressed during the Giardia life cycle. Next, we explored whether genetic exchange occurs between nuclei during the Giardia life cycle. In a preliminary account of these experiments, we reported nuclear fusion and exchange of episomes during encystation (which we termed “diplomixis”), as well as the cyst-specific expression of several HMGs (Poxleitner et al., 2008). I have since built upon these results, expanding my analysis to include all seven HMGs and obtaining more precise HMG expression patterns using endogenous tagging. In addition, I have now directly demonstrated that chromosomal genetic exchange occurs between nuclei in the cyst. I report those results here, as well as relevant data from the published paper.

Results Three HMGs are expressed only during encystation To determine when and where the HMGs are expressed during the Giardia life cycle, we used plasmid integration (Gourguechon and Cande, 2011) to endogenously tag all seven HMGs with epitope tags. Because the genes were endogenously tagged, the constructs were under the control of the native promoter. We then examined the expression and localization of these proteins in both trophozoites and cysts. Mer3 was not localized to the nuclei of trophozoites or cysts (data not shown), suggesting that it may not, in fact, be a Mer3 ortholog; 72 consistently, it is annotated as an RNA helicase in the Giardia genome database (ORF GL50803_87022 at giardiadb.org). As shown in Figure 4.1, only Dmc1b was expressed at high levels in the trophozoite; Mnd1 and Hop2 were present at low levels, and the other HMGs (Spo11, Dmc1a, Hop1) were absent. In contrast, Dmc1b was absent in cysts, while Spo11, Dmc1a, and Hop1 were dramatically upregulated, appearing in encysting trophozoites and remaining in fully encysted cells [Figure 4.1 and (Poxleitner et al., 2008)]. Mnd1 and Hop2 remained at low (but detectable) levels. In all cases and as expected, the proteins were localized to the nuclei in a punctate pattern.

Genetic exchange occurs between nuclei in the cyst Based on the timing of expression of Spo11, Dmc1a, and Hop1, we decided to focus on encystation as a time for potential genetic exchange. As previously reported, the nuclei are able to fuse and exchange episomal plasmids during encystation, a process we have termed “diplomixis” (Poxleitner et al., 2008). We also observe increased staining of encysting cells by terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay (Figure 4.2), further suggesting that homologous recombination could be occurring during diplomixis. To test this hypothesis, we integrated a plasmid containing a 10-kb array of LacO repeats. To ensure the presence of only the integrated construct, we made clonal populations and tested for the absence of episomes by PCR (Figure 4.3). Like episomes (Yu et al., 2002), integrated plasmids are found in only one of the two nuclei in the trophozoite (Figure 4.4A). We then performed fluorescence in situ hybridization (FISH) using a fluorescent probe to label the LacO repeat. In the starting population, 76% of cells (n = 100) contained one spot, while 24% of cells contained no spot (Figure 4.4A). Because this population was clonal and the marker was integrated (and therefore stable even without drug selection), we interpret this result as an artifact of FISH; however, it remains possible that the cells lost the repeat cassette via recombination. After encystation, excystation, and growth of excysted trophozoites to confluency, we observed that 47% of cells (n = 100) had one spot, 31% had no spots, and 22% had two spots (one in each nucleus) (Figure 4.4A). The observed pattern could be generated in two ways: 1) genetic exchange between nuclei, or 2) nuclear sorting such that during excystation, each daughter cell received two identical nuclei. However, because the majority of cells contained one labeled nucleus and one unlabeled nucleus, nuclear sorting would have had to occur in a minority of excysting cells (excyzoites). To begin to distinguish between these possibilities, we also performed FISH on cysts and excyzoites (Figure 4.4B). As expected, the nuclei of most cysts contained two clustered pairs of nuclei, each with one labeled nucleus. This pattern was recapitulated in excyzoites, again supporting the idea that the non-identical nuclear pairs remain associated throughout encystation and excystation. While we were able to observe an occasional cyst with three labeled nuclei (second panel in Figure 4.4B), this was rare; however, we did notice that the spots in the cyst were often positioned between the nuclei (first panel in Figure 4.4B),

73 suggesting that the nuclei might remain fused in the cyst and only separate during excystation. To further distinguish between genetic exchange vs. nuclear sorting during excystation, we examined a strain that contained the same LacO repeat construct, but also contained a TetO repeat construct integrated into a different chromosome in the same nucleus (Figure 4.4C). As with the LacO construct, the absence of episomal plasmid in this clonal line was confirmed by PCR. As observed in the single-marker strain, each nuclear pair in the pre-encysting trophozoites contained both markers in a single nucleus. In excysted trophozoites, however, we observed trophozoites with all possible arrangements of markers (one in each nucleus, two in one nucleus and one in the other, and two in both nuclei), further indicating that genetic exchange occurs between nuclei during encystation.

Cyst nuclei display dynamic movements For homologous recombination to take place between nuclei in the cyst, the nuclei must come together and fuse. As we previously reported, transmission electron microscopy of cysts reveals fused nuclei with continuous nuclear envelopes (Poxleitner et al., 2008). Importantly, we never observe spindles in cysts, so what is the driving force for this fusion (and the resulting fission that must occur prior to excystation)? To address this question, we collected time-lapse images of live cysts stained with acridine orange to label the nuclei. We discovered that the cyst nuclei display dynamic behaviors (Figure 4.5A-E), moving as pairs within the cyst. As shown in Figure 4.5, the pair on the right moves relative to the pair on the left, and in ~10 seconds rotates from a vertical (Figure 4.5A) to a horizontal (Figure 4.5C) orientation. The nuclei within the pair are closely opposed such that they often appear as a single mass. The pair on the left also moves as a single unit (Figure 4.5F-J). The nuclear movements in cysts are very different from what is observed in trophozoites, in which the two nuclei are tethered to the cytoskeleton and exhibit no movement except during mitosis (Poxleitner et al., 2008). As demonstrated in Chapter 3, the cyst nuclei are closely associated with bundles of internal axonemes, and these axonemes also continue moving in the cyst (Figure 4.6). Thus, we speculate that the nuclear movements are the result of this residual flagellar movement. These dynamic movements may provide the force necessary for karyogamy to occur and promote the mixing of nuclear contents.

Discussion On the basis of these data we propose a model for diplomixis during encystation (Fig. 4.7). First, at the induction of encystation, the nuclei divide with two separate spindles (see Chapter 3). This division generates a cyst with two pairs of non-daughter nuclei at opposite ends of the cell. Each pair of nuclei appears to be associated with a bundle of internal axonemes and moves together within the cyst, perhaps due to the beating of these axonemes. One pair of nuclei then migrates to the opposite end of the cell to join the other nuclear pair. At some point, karyogamy 74 between two non-daughter nuclei can occur, resulting in the appearance of three nuclei (Poxleitner et al., 2008). This three-nuclear stage is commonly observed with all nuclei in close proximity to one another, so the actual fusion probably occurs after the nuclei have migrated together at one end of the cell. Finally, because fully mature cysts have four nuclei by the initiation of excystation, we deduce that nuclear fission and the reformation of four separate nuclei must follow karyogamy and genetic exchange. This parasexual process, which we call diplomixis, is thus far unique to Giardia, although we predict it occurs in other members of the order Diplomonadida. Unlike automixis, diplomixis is not accompanied by a meiotic genome reduction and the subsequent fusion of gametes from the same parent; furthermore, unlike the parasexual cycle of some fungi, diplomixis does not appear to involve the fusion of two individual cells (Archetti, 2010), at least in vitro. Importantly, our results here support our previous observations that during excystation, the daughter trophozoites receive one copy of each parental nucleus. The distribution of identical nuclei to daughter cells during excystation would explain the low levels of heterozygosity observed in Giardia populations. Thus, our observation that nuclei remain in non-identical pairs in the cyst and excyzoite and that the majority of excyzoites contain non-identical nuclei (Figure 4.4) is significant and implies that there are other mechanisms in place to reduce genomic heterozygosity in Giardia. Homologous recombination between nuclei, if accompanied by mitotic recombination/gene conversion within nuclei, could be one of those mechanisms. It remains possible that a minority of cells undergo nuclear sorting, and the patterns of markers shown in Figure 4.4C are the result of failure of the FISH probe to label all of the markers present in the cell. However, based on our observations of the tight association between nuclear pairs throughout encystation and excystation, we find this possibility unlikely. Based on our results using integrated markers, we currently cannot distinguish between homologous recombination and the exchange of whole chromosomes during diplomixis. However, the timing of expression of the HMGs and the increase in TUNEL staining during encystation all suggest that homologous recombination is occurring. Regardless of the mechanism, genetic exchange appears to be occurring at relatively high frequencies, as we observed 22% of excysted cells with both markers (Figure 4.4A). Indeed, this is likely an underestimate of the amount of exchange occurring, as our assay can only detect events involving transfer of the marker. This finding is consistent with diplomixis being used as a mechanism to reduce nuclear heterozygosity: to be effective, genetic exchange would be expected to be relatively common and involve large portions of the genome. Interestingly, in the strain with two integrated markers, we were able to observe cells with one marker in each nucleus. Because of the tight association of the nuclei in the cyst, the simplest interpretation of this pattern is that a marker moved from one nucleus in the pair to the other. However, during meiotic homologous recombination, only one sister chromatid is involved, and so the other

75 sister should be unaffected by recombination and retain the marker. Further investigation will be required to determine whether recombination, if it occurs, takes place before or after S phase in the cyst (which increases from 8N in the encysting trophozoite to 16N after cyst formation (Bernander et al., 2001)). We also occasionally observe two paired spots rather than one in some cells (see fourth panel in Figure 4.4). These spots could represent replicated chromosomes that became separated; however, it is also possible that these patterns result from recombination and gene conversion between homologous chromosomes in the same nucleus. Again, further investigation will be necessary to distinguish between these possibilities. The observation that Spo11, Hop1, and Dmc1a are only expressed during encystation suggests that homologous recombination is occurring during diplomixis. If this turns out to be the case, it would be one of the first examples of these proteins functioning in a non-meiotic context. Recently, Spo11 has been found to mediate homologous recombination during the parasexual cycle of the fungus Candida albicans (Forche et al., 2008). However, the lack of a sexual cycle in this organism is clearly a case of loss, whereas it is unclear whether Giardia or its relatives have a true meiotic cycle. Thus, future studies of the functions of the HMGs during diplomixis in Giardia could shed light on the evolution of the meiotic homologous recombination machinery. Our finding that Dmc1b is expressed only in the trophozoite suggests that, despite its annotation as a Dmc1 homolog based on phylogenetic analysis, this protein actually has a Rad51-like function in DNA damage repair. The absence of this protein in the Giardia cyst suggests that either it is still involved in diplomixis but present at undetectable levels, or somatic homologous recombination and diplomixis are compartmentalized in Giardia, with different machinery used for each process. This would be different from the case in most sexual organisms, in which Rad51 and Dmc1 work together to facilitate meiotic homologous recombination (Masson and West, 2001). However, a recent study found a similar specialization of Rad51 and Dmc1 during meiosis in Tetrahymena, in which Dmc1 appears to be the primary player in meiotic homologous recombination (Howard-Till et al., 2011). While diplomixis could explain the maintenance of low heterozygosity between nuclei, it is also possible that rare meiotic events occur in the wild, as suggested by several recent population genetics studies (Cooper et al., 2007; Lasek- Nesselquist et al., 2008). These studies found evidence for recombination within and between assemblages, which cannot be explained by diplomixis within a single cell. Cell fusion during encystation followed by diplomixis could produce patterns consistent with these findings; however, as shown in Chapter 3, cysts are primarily derived from a single cell, at least in vitro. Future studies should aim to test for fusion or a true meiotic event in vivo, similar to what has been done to detect sexual events in parasites like Trypanosomes and Leishmania (Akopyants et al., 2009; Peacock et al., 2011). Identifying this process may be difficult, particularly if it is rare or occurs only under certain conditions. Nevertheless, a deeper understanding

76 of sexual processes in Giardia may shed light on the evolution of meiosis and give us deeper insight into the population genetics of this important parasite.

Materials and Methods Culture conditions Giardia intestinalis trophozoites, WB clone C6 (ATCC 50803), were grown in modified TYI-S-33 medium (Keister, 1983) with bovine bile (Sigma-Aldrich, St. Louis, MO). Cultures were incubated in 16-mL plastic screw top tubes (Fisher Scientific, Pittsburgh, PA) at 37°C and passaged every three days.

In vitro encystation Encystation was performed essentially as described (Kane et al., 1991). Briefly, trophozoites were seeded into 30- or 50-mL tissue culture flasks (BD Biosciences, Bedford, MA) and grown to 80%-100% confluency. The media was then decanted and replaced with prewarmed encystation media (10 mg/mL bovine/ovine bile, pH 7.8). After 24 hours at 37°C, cells were either fixed for observation or incubated on ice for 15 min, pelleted at 500 xg, and resuspended in regular growth medium (0.5 mg/mL bile, pH 7.1). After an additional 24 hours of growth at 37°C, the cells were again incubated on ice, pelleted, and resuspended in 50 mL ice-cold, sterile deionized water to destroy all unencysted cells. Cysts were stored at 4°C for 1-3 days, then excysted as described below.

In vitro excystation Excystation was performed essentially as described (Boucher and Gillin, 1990). Briefly, water-treated cysts were pelleted in a 50-mL conical tube, then resuspended in 10 mL prewarmed stage I solution (1X Hank’s Balanced Salt Solution [Invitrogen, Carlsbad, CA], 57 mM L-cysteine HCl, 32.5 mM reduced glutathione, and 25mM NaHC03; the pH was adjusted to 4.0 with 0.01N HCl). The tube was vortexed briefly, then incubated at 37°C for 30 minutes. After another brief vortexing, cells were pelleted at 500 x g for 5 minutes. The supernatant was then decanted and replaced with 10 mL prewarmed stage II solution (Tyrode’s buffer [Sigma-Aldrich, St. Louis, MO] containing 1 mg/mL Trypsin II-S [Sigma]; the pH was adjusted to 8.0 with 1 M NaHC03). The tube was vortexed briefly, incubated at 37°C for 30 minutes, then vortexed again and pelleted. The supernatant was decanted and replaced with prewarmed growth media (see Culture conditions), and cells were moved to 16-mL culture tubes and incubated at 37°C. Cells were then either fixed 1-3 hours later for immunofluorescence staining or grown to confluency (approximately 5-7 days).

Immunostaining Fixation and immunostaining were performed essentially as described (Carpenter and Cande, 2009). Briefly, cells were fixed in 1X PEM buffer (100 mM PIPES, 1 mM 77 EGTA, 0.1 mM MgSO4) containing 2% paraformaldehyde. After fixation for 30 minutes at 37°C, cells were washed once with PEM buffer, then resuspended in PEM buffer and attached to poly-L-lysine-coated coverslips for 30 minutes. Alternatively, for delta-giardin antibody staining, cells were fixed for 3 minutes in ice-cold methanol, then washed twice in PBS before being attached to poly-L-lysine coverslips. Fixed cells were permeabilized with 0.1% Triton-X-100 for 15 minutes, washed three times with PEM, and blocked in PEMBALG (PEM plus 1% BSA, 0.1% NaN3, 100 mM lysine and 0.5% gelatin from cold water fish skin [Sigma, ]) for 2 hours. Primary antibody incubations were conducted at room temperature overnight, and antibodies were diluted in PEMBALG as follows: mouse anti-HA antibody 1:100 (Sigma); goat anti-mouse 488 secondary antibody 1:200 (Invitrogen)) at room temperature for two hours. After a final three washes each in PEM and PEMBALG, coverslips were mounted onto slides with Prolong Gold Antifade Solution with DAPI (Invitrogen).

Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) TUNEL staining was performed using an In Situ Cell Death Detection Kit according to the manufacturer’s instructions (Roche, Basel, Switzerland).

Creation of clonal strains containing integrated LacO and TetO constructs For the LacO array integration construct, a SacI/PvuII fragment containing part of the Spo11 coding sequence and a puromycin resistance cassette (from the vector pKS-Spo11-3HA-pac) was inserted into the LacO array-containing plasmid SR7 ((Rohner et al., 2008), a gift from S. Gasser) cut with restriction enzymes SacI and EcoRV (Fig. 4.3). For integration, the plasmid was linearized with MfeI, which cuts within the sequence for Spo11. After the selection of stable transformants, single- cell clones were obtained by limiting dilution as previously described (Baum et al., 1988). These clones were screened for the presence of the integrated plasmid and absence of episomal plasmid by PCR as shown in Figure 4.3, using the primers P1 (5’- CATGATTTTCTTGTTTTAGGG -3’) and P2 (5’- TAATACGACTCACTATAGGG- 3’) or P1 and P3 (5’- AGACTCTACAGGATACTTTCAG-3’). PCRs were performed by boiling a concentrated culture (200 µl) of approximately 1.3 x 107 cells for 10 minutes at 95°C. One microliter (~65,000 cells) was then used for PCR amplification.

For the TetO array construct, an EcoRI/XbaI fragment containing the median body protein (MBP) coding sequence and a neomycin resistance cassette (from the vector pKS-MBP-neo) was inserted into the TetO array-containing plasmid SR8 ((Rohner et al., 2008), a gift from S. Gasser) cut with the restriction enzymes EcoRI and NheI. For integration, the plasmid was linearized with MfeI, which cuts within the sequence for MBP. After the selection of stable transformants, clonal lines were created and confirmed as described above, except using primer P4 (5’-TTC CGA ATT TTG AGA CAC CTC-3’) instead of primer P3 listed above. 78 Fluorescence in situ hybridization FISH was performed essentially as described (Poxleitner et al., 2008). However, instead of using nick translation to create the probe, a Cy3-labeled LacO oligonucleotide probe (5’-CCACAAATTGTTATCCGCTCACAATTCCACATGTGG-3’) and/or a FITC-labeled TetO oligonucleotide probe (5’- TCGAGTTTACCACTCCCTATCAGTGATAGAGAAAAGTGAAAG -3’) was used at a concentration of 5 ng/µl.

Fluorescence deconvolution microscopy Images were collected with SoftWorX image acquisition software (Applied Precision, Issaquah, WA) on an Olympus IX70 wide-field inverted fluorescence microscope with an Olympus UPlanApo 100X, NA 1.35, oil-immersion objective and Photometrics CCD CH350 camera cooled to -35°C (Roper Scientific, Tuscon, AZ). Serial sections were acquired at 0.2-µm intervals, and data stacks were deconvolved using the SoftWorX deconvolution software. For presentation purposes, two- dimensional maximum intensity projections were created from the 3D data sets using the DeltaVision image analysis software (Applied Precision).

Live cell staining and visualization Cells were induced to encyst for 21-24 hours and then collected by centrifugation. For acridine orange staining, one milliliter of cells was stained to a final concentration of 10 µM acridine orange (Invitrogen). For GFP-tubulin visualization, encysting cells expressing GFP-tagged beta tubulin (a gift from S. Dawson) were incubated in HEPES-buffered saline for 20 minutes to facilitate GFP fluorescence prior to viewing. Cells were observed on slides with a Zeiss Axiovert 200M wide- field inverted fluorescence microscope with a 63x PlanApo, NA 1.4, oil-immersion objective. Time-lapse images were collected every second with Metamorph software (Molecular Devices, Downingtown, PA).

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82 Figure 4.1: Expression of the meiosis-specific gene homologs in trophozoites and cysts. Epitope tags were integrated into the endogenous loci of the six giardial meiotic gene homologs, creating C-terminally tagged proteins under their native promoters. Control (wild-type) and tagged strains before and after encystation were labeled by immunofluorescence using an anti-HA antibody (green) and stained with DAPI to label nuclei (blue).

83 Trophozoite Cyst DNA α-HA Merge DNA α-HA Merge

WT

Spo11

Hop1

Hop2

Mnd1

Dmc1a

Dmc1b

84 Figure 4.2: TUNEL staining increases during encystation. (A) Terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) was performed to detect double-strand breaks in untreated trophozoites (negative control), DNase treated trophs (positive control) and methyl methanesulfonate (MMS)-treated trophs (positive control). TUNEL staining is shown in green and DAPI-stained nuclei are shown in blue. (B) Encysting cells were stained with TUNEL and immunolabeled with a cyst wall protein (CWP) antibody (red), which labels encystation-specific vesicles in the encysting trophozoite and the cyst wall in the mature cyst. TUNEL staining is shown in green and DAPI-stained nuclei are shown in blue.

85 A DNA TUNEL Merge

Negative control

Positive control (DNase treated)

0.01% MMS

Merge B DNA TUNEL w/CWP

Encysting trophozoite

Cyst

86 Figure 4.3: Scheme for the creation and verification of a clonal strain with an integrated marker. (A) Map of the plasmid used for integration, which contains an array of 256 LacO repeats. The binding sites for primers P1 and P2 are shown, as is the restriction site (MfeI) that was used to linearize the plasmid for integration. (B) Schematic of the linearized plasmid, showing the binding sites for primers P1, P2, and P3. Primers P3 (which binds upstream of where the plasmid is integrated in the genome) and P1 were used confirm integration of the construct, and primers P1 and P2 (which should not produce a product in the absence of circular plasmid) were used to check for the absence of the circular plasmid. (C) Results of PCR testing of the wild-type (WT), parental (non-clonal) transformed strain, and clonal strain used in subsequent experiments. The presence of a plasmid-specific product in the parental strain suggests that either residual episome was present or that some cells integrated the plasmid in tandem arrays, which would also produce a product with primers P1 and P2.

87 A Plasmid

P2

P1

B P3 Integrated P2

P1 chromosome chromosome

C Primers P1 and P2 Primers P1 and P3

WT Parent CloneWT Parent Clone

88 Figure 4.4: Nuclear identity and genetic exchange during Giardia differentiation. (A) A clonal population of cells containing an integrated LacO array in one nucleus were subjected to fluorescence in situ hybridization (FISH) to track the marker in the starting population (pre-encystation trophozoites) and in excysted trophozoites. DAPI-stained DNA is shown in blue and the LacO probe is shown in red. Percentages of cells with 0, 1, and 2 spots are shown below each image (n = 100). (B) Two cysts and an excyzoite from the experiment described in (A). Cyst wall protein (CWP) staining is shown in green; background staining of the excyzoite is shown to outline the cell shape. DAPI-stained DNA is shown in blue and the LacO probe is shown in red. Note the presence of one spot in each nuclear pair in the first cyst and in the excyzoite and the presence of three spots in the second cyst. (C) Patterns observed after excystation of a clonal population containing two markers, LacO (red) and TetO (green), integrated in the same nucleus (first panel). The number of nuclei labeled and the number of spots are indicated below each image (e.g., 1N: 1R 1G = one nucleus, one red spot and one green spot). All scale bars = 5 µm.

89 A One nucleus Two nuclei No signal

Pre-encyst population 76% 0% 24% Excysted population 47% 22% 31%

B

Cyst Cyst Excyzoite

C

1N: 1R 1G 2N: 1R 1G 2N: 1R 2G 2N: 2R 1G 2N: 2R 2G

90 Figure 4.5: Dynamic nuclear movements in the Giardia cyst. Selected frames five seconds apart from a movie of a live, acridine orange-stained cyst. The two pairs of nuclei are indicated with asterisks in (A). The arrowhead follows the movement of the nuclear pair on the right side of the cyst (A-E), and the arrow follows the movement of the pair on the left (F-J). Note the close apposition of the two nuclei on the right.

91 A B C D E * *

F G H I J

92 Figure 4.6: Beating of internal axonemes in the Giardia cyst. Selected frames five seconds apart from a movie of a live cyst expressing GFP- tagged beta tubulin. A disc fragment (asterisk) and two bundles of internal axonemes are visible. The arrowhead follows the movement of one bundle of axonemes as it beats within the cyst.

93 A B C D E *

94 Figure 4.7: Model of diplomixis. A trophozoite has two independently inherited nuclei, blue or red, and only one (red) contains an integrated marker. Of the meiosis-specific gene homologs, only Dmc1b, Mnd1, and Hop2 are expressed at this stage. During encystation, Dmc1a, Spo11, and Hop1 begin to be expressed, and each nucleus divides, resulting in two pairs of non-daughter nuclei at opposite ends of the cell. After migration of the nuclei to one pole of the cell, fusion can occur between non-daughter nuclei (i.e., between a red and a blue nucleus), leading to the exchange of genetic material, possibly facilitated by the meiotic genes. Subsequently, the fused nuclei undergo fission and move apart, resulting in three (red) nuclei containing the integrated marker. The nuclear pairs remain associated during excystation, producing trophozoites with either one red and one blue nucleus or two red nuclei.

95 Excystation

No exchange Encystation

Dmc1a Dmc1b Spo11 Exchange Mnd1 Hop1 Hop2 Mnd1 Hop2

96