Featured article #6

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How to Cheat Death: Genes Involved in Reverse Development

INTRODUCTION

The regeneration or rejuvenation of animal cells has always been an interesting topic and represent an actual field of challenge and debate due to the many potential applications in the medical field. Regeneration can possibly arise by transdifferentiation of existing cells or by stem cells (totipotent cells that can generate any differentiated cell). In both cases, the response occurs according to chemical, genetic, and physiological signals. These mechanisms can be activated to give rise to a whole array of new cell types (Brockes, 1998; Slack, 2003). A deeper understanding of the mechanisms underlying rejuvenation would provide a first step towards interpreting the biology of senescence and bringing new insight for future application on regenerative medicine.
Cnidarians are incredibly diverse animals exclusively found in aquatic and mostly marine environments. Jellyfish, corals, sea anemones, and hydras are among the most prominent representatives of this taxon. One of their typical features is the presence of cnidocytes, specialized cells that are used for prey capture and defense. Cnidarians have long been considered simple animals but they present a variety of complex life cycles and developmental patterns. In addition, they are the only known group of organisms with the potential to undergo reverse development (RD) (Schmich et al., 2007). Reverse development was first discovered in scyphozoans more than a century ago (Hadzi, 1909) and represents an unparalleled feature of cnidarians within the animal kingdom (Piraino et al., 2004). Since the discovery of this mechanism, several hydrozoan species have shown the potential to reverse their ontogenetic programs (Piraino et al., 2004).
Under normal conditions, the hydrozoan general life cycle is characterized by the alternation of a post-larval benthic polyp and an adult pelagic medusa (Figure 1). The sexually mature adult medusa releases gametes, which, upon fertilization, form short-lived lecithotrophic (non-feeding) planulae. The planulae settle on to a substrate and form larval hydroids. The hydroids grow asexually, usually forming colonies. Through a complex process that involves the entocodon (medusary nodule), hydroids produce and release medusae, completing the cycle (Boero & Bouillon, 1987; Boero et al., 1998; Boero et al., 2002).Interestingly, some hydrozoan species are also able to revert their life cycle. In these cases, the adult medusa goes back to the juvenile stage of polyp as an adaptive strategy to deal with environmental stress (Carlà et al., 2003; Piraino et al., 2004; Schmich et al., 2007).

To date, different cnidarian species have been identified as performing RD, such as Turritopsis nutricula, T. dohrnii (Oceanidae), Laodicea undulate (Laodiceidae), and Hydractinia carnea (Hidractiniidae; synonymous of Podocoryne carnea, [Schuchert, 2011]) (Piraino et al., 2004; De Vito et al., 2006; Schmich et al., 2007). Furthermore, species belonging to the Turritopsis genus are among the few known species with the ability to perform RD from a free-living medusa to a polyp state after becoming sexually mature (Bavestrello et al., 1992; Piraino et al., 1996). Through stereoscopy observation, Carlà and colleagues (2003) identified four stages during the life cycle of T. nutricula and categorized them as healthy medusa, unhealthy medusa, four-leaf clover and cyst. According to these authors, the healthy medusa has a bell-shaped umbrella with long tentacles and swims actively. The unhealthy medusa is not able to swim and maintains its tentacles in a retracted position, and the typical transparency of the normal healthy medusa is lost. The four-leafed stage is characterized by the absence of tentacles and the reduction of the sub-umbrella cavity, which shows a number of lobes and many degenerative processes. During the cyst stage the organism is spherically shaped, with a smooth surface; this cyst is able to attach to the substrate and rapidly give rise to a polyp stage. Although RD was identified in several species, most of them undergo some restrictions in expressing transdifferentiation at the adult stage. For instance, in Hydractinia carnea, the potential for RD is lost in older medusa stages (Schmich et al., 2007).

Reverse development (or cell transdifferentiation) is a process through which a mature somatic cell transforms into another mature somatic cell without undergoing an intermediate pluripotent state or progenitor cell type (Graf & Enver, 2009). Transdifferentiation also involves a change in commitment and gene expression of well- differentiated, non-cycling somatic cells to other cell types (Okada, 1991). In some medusa species, this process occurs in all cells, e.g. those of the manubrium, tentacular bulbs, radial canals, exumbrellar rim, marginal canal, subumbrellar rim, or even gonads (Piraino et al., 2004). Degenerative and apoptotic cellular morphological modifications are involved during the RD of T. nutricula. It has been shown that the gonads are retained throughout all stages, thus showing evidence that both young and mature medusa can undergo reversion (Carlà et al., 2003).

The advent of molecular and genomic technologies has provided an available toolkit for experimental studies on cnidarians development. It is now possible to manipulate and analyze tissues and embryos according to several approaches, such as mRNA injection, RNAi, and transgenesis (Technau & Steele, 2011). Transdifferentiation is a very interesting mechanism that can provide many insights for the development of tolls and products for regenerative medicine, such as treatments for heart diseases. Postnatal cardiomyocytes have little or non-regenerative capacity; although, endogenous cardiac fibroblasts are able to differentiate in beating cardiac cells replacing damaged cardiomyocytes. In a study performed by Ieda (2010), a specific combination of three transcription factors was identified as being able to generate functional beating cardiomyocytes directly from mouse postnatal cardiac fibroblast. In addition, the induced cardiomyocytes were globally reprogrammed to a cardiomyocytes-like gene expression profile.

The main goal of this proposal is to investigate which signaling networks are involved in stage differentiation during hydrozoans life cycle, in order to identify and understand the mechanisms underlying reverse development. We will thus focus on three marine hydrozoans, Turritopsis dohrnii, Hydractinia echinata, and Hydractinia carnea as model organisms for reverse ontogeny and dedifferentiation of somatic cells to multipotency. Turritopsis dohrnii and Hydractinia carnea were previously compared under different chemical and physical conditions and showed a different potential for RD (Schmich et al., 2007). To date, there are no evidences of RD in Hydractinia echinata. However, its transcriptome was completely sequenced and the data is available at http://www.mchips.org/hydractinia_echinata.html. Since H. echinata is a sister group of H. carnea, which has the ability to perform transdifferentiation, we will also compare both transcriptomes in an attempt to identify differentially expressed genes.

We strongly believe that our results will pave the way for future studies aiming to clarify the mechanisms of senescence and rejuvenation.


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Figure 1. Models of some hydrozoan life cycles (removed from Piraino et al., 2004). (A) Hydra spp. has no medusa stage and no planula larva. Asexual budding of new polyps is the most frequent mode of reproduction. (B) Hydractinia echinata has a highly reduced medusa stage that remains attached to the polyp in the form of fixed gonophores. Fertilization leads to the formation of a planula larva, which will undergo metamorphosis to develop into a polyp colony. (C) The life cycle of Hydractinia (Podocoryna) carnea includes a swimming medusa stage and a planula larva. Artificial detachment of late medusa buds leads to the development of medusae that are complete but reduced in size. In contrast, artificially detached early medusa buds are capable of transformation back into polyps. Such reverse transformation is not usually achieved by late medusa buds or liberated medusae. (D) Turritopsis nutricula has a typical three-stage life cycle: planula, polyp, and medusa. However, medusae at all stages of development retain the potential for life-cycle reversal; even spent medusae do not die, but transform back spontaneously into new polyp colonies (Piraino et al., 2004).


RESEARCH DESIGN AND EXPERIMENTAL METHODS

Sampling, Culture and Stage selection for RNA extraction

Samples of T. dohrnii and H. carnea will be collected from the Mediterranean Sea and Naples, respectively. The specimens, separated by species and sampling site, will be maintained in two different aquariums, one with filtered sea water and sediment from the sampling location and the other with artificial sea water and autoclaved sediment, both on 22°C; all the aquariums will be equipped with aeration system and a moderate continuous water flux. The cnidarians will be fed with Artemia naupli and will be monitored to check the developmental stage until the first life cycle is complete.
RNA will be extracted from T. dohrnii and H. carnea at different developmental stages: medusa buds, older medusa stage, planula, ball-like stage, stolon, polyp formation, and adult polyp (Figure 2). Life stages will be selected under binocular microscope (Schmich et al. 2007) and pictures will be taken through a camera coupled to the microscope.

To induce RD, two pilot experiments will be performed using heat shock and CsCl incubation. The more effective protocol will be select to induce RD of medusa in both species. During the RD, eight life stages will be select for RNA extraction (Figure 2). Subsequently, the extracted RNA samples will be used for library construction. All bioassays will be done in duplicates to confirm the obtained results.

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Figure 2. Selected life stages during reverse development for RNA extraction. The description of each state is provided.

Transcriptomic analyses

The transcriptomes of Turritopsis dohrnii and Hydractinia carnea will be analyzed through a comparative approach in order to identify genes and signaling pathways that are related to reverse development. Genes differentially expressed during RD will be selected for experimental validation using RNAi (see below for details). Subsequently, a Blast search will be performed to retrieve potential homologs in the Hydractinia echinata transcriptome data. Since this species has medusa stages morphologically reduced and remaining attached to the polyp, it is not known if genes involved RD are present and activated in H. echinata (Figure 3).

Group_Assignment_SPSAS_Evo_280812_Almost%20done.jpg

Figure 3. Hydractinia echinata and Podocoryne carnea (Syn. Hydractinia carnea): life cycle. Taken from Plickert et al., 2012.

mRNA-Seq library preparation

A pool of 10 individuals from each developmental stage (Figure 2) from T. dohrnii and H. carnea will be isolated for RNA extraction with Trizol. Total RNA (5 μg) will be used to prepare the transcriptome library using the mRNA-Seq preparation kit (Illumina) following the manufacturers instructions. The mRNA will be purified using magnetic oligo-dT beads and fragmented. First strand synthesis of cDNA will be done using SuperscriptII reverse transcriptase followed by RNase H treatment. Second strand cDNA synthesis will be performed using DNAPol1. Paired end adaptors (Illumina) will be ligated to 5′ and 3′ ends of DNA fragments followed by PCR amplification using adaptor specific primers. PCR amplified products will be gel purified, quantitated by Nanodrop and Bioanalyzer and sequenced on an Illumina GAIIx or HiSeq2000 platform. We will use Illumina’s version 4 or TruSeq sequencing chemistry. Each library will be loaded in one lane of a Single Read v4 or HiSeq flow cell (Illumina) at a concentration of 6 pM. All libraries will be sequenced to 50– 51 cycles in one direction.

Assembly and annotation

To eliminate low quality nucleotides, raw Illumina reads will be trimmed using a custom Perl script with windowed adaptive trimming (Phred quality threshold of 20 and minimum read length of 20 nt). The trimmed reads will be separately fed into Rnnotator, automated de novo RNAseq assembly pipeline (Martin et al., 2010), to remove duplicates and erroneous sequences. Multiple rounds on velvet assemblies will then be performed using different hash values to account for the different sequencing depth for different transcripts (Martin et al., 2010). Resulting contigs will be merged using Minumus2 from the AMOS package (Martin et al., 2010; Sommer et al., 2007). The resulting contigs and remaining singletons will be aligned with BLASTX to the complete UniProt database (www.uniprot.org/downloads). Acceptable hits will be determined by a bitscore of >45 and a corresponding e-value of <1−5. To maximize the information content, obtained results will be improved with lower ranking (but more informative) annotations using a PERL script and a list of keywords to avoid (i.e. ‘uncharacterized protein’ and ‘predicted protein’). Contig and singleton sequences will be also submitted to the KEGG automated annotation server (www.genome.ad.jp/tools/kaas/) for further functional annotation. Pfam and Smart domains will be mapped to assembled transcripts using RPS-BLAST with an e-value threshold of 1e−5. Gene Ontology (GO) annotations will be used to assign biological functions to genes included in this study (Ashburner et al., 2000). GO annotations will be transferred from Pfam and Smart annotations mapped to our transcripts.

Experimental Validation of Predicted Transcripts

First strand cDNA will be synthesized from 1 μg of total RNA using SuperScriptII (Invitrogen). PCR amplification with specific primers will be performed for 20, 25 and 30 cycles and the samples resolved on 1.2% agarose stained with ethidium bromide. Experimental validation of transcript models and predicted patterns of expression will be performed using semi-quantitative RT-PCR. We will select a total of 20-30 transcripts as candidates for RT-PCR and construct PCR primers against their sequences. Transcripts predicted as being differentially expressed during RD or alternatively spliced will be selected chosen as candidates for validation, and when appropriate, we will choose candidates highly expressed or biologically interesting. Validation studies will be designed to test the expression of (1) transcript assemblies, (2) differential expression, and (3) alternative splicing.

We will assess the significance of differential gene expression with edgeR (Ewen- Campen et al., 2011), version 2.0.5 according to standard protocols outlined in the package manual. These analyses will be run in R version 2.12.2

Functional analysis of candidate genes by knockdown using RNAi technique

Preparations of double-stranded RNA (dsRNA) will be performed as described before (Lynch & Desplan, 2006) with minor modifications. Fertilized medusas will be placed in individual wells in a 24-well plate and the incubated with dsRNA (100μl of sterile seawater with dsRNA 8ng/μl) (Wittig et al., 2011). As control, a group of medusas will be treated with a non-related dsRNA for 72 hours, then, metamorphosis will be induced by adding 580mM CsCl (1:5 seawater) for 3 hours. To finish the dsRNA treatment, the sample must be washed 3 times with seawater. At last, animals will be fixed in paraformaldehyde 4%.The effects of dsRNA treatment will be observed through microscope and registered though a camera coupled to the microscope.


EXPECTED RESULTS

The present proposal expects to obtain the transcriptome of two different species, in many developmental stages. Taking advantage of a comparative approach within and between species, will allow us to identify the gene networks involved in the transdifferentiation process. Finally, we expect to experimentally confirm the function performed by these genes involved in reverse development as well provide guidelines for future contributions to medical research.


FUTURE DIRECTIONS

Different species of marine hydrozoans display a unique potential to reverse their life cycle (Piraino et al., 2004; De Vito et al., 2006; Schmich et al., 2007). It is believed that this backward ontogeny is coordinated through activation of specific genetic switches leading to transdifferentiation and body plan reorganization (De Vito et al., 2006). However, the genetic mechanisms underlying this process are still unknown. Our proposal aims to improve the understanding in this field, providing a solid basis towards reverse development process and the genes involved on it.

Preliminary attempts to trace the evolutionary relationships between cnidarian and vertebrate stem cells centered on searching sequenced cnidarian genomes and EST datasets for homologs of the four pluripotent genes known to be expressed in vertebrate stem cells (Klf4, Oct4, Sox2 and Nanog). However, homologs of these genes have not been unambiguously identified in the Nematostella or Hydra genome (Chapman et al., 2010), suggesting that either the role of these key genes is carried out by other related genes or instead that stem cells machinery evolved independently in cnidarians and vertebrates (Technau & Steele, 2011).

Several genes and signaling pathways associated with patterning and developmental processes in bilaterians, including those related with human diseases, are also present in cnidarians (Sullivan & Finnerty, 2007; Sullivan et al., 2008; Soza-Ried et al., 2009).The combination of the unique characteristics of these organisms coupled with their phylogenetic position, make them ideal models to investigate the evolution of many key aspects of animal development, such as the formation of the third germ layer, the nervous system, the generation of bilaterality, the ability to undergo rejuvenation, aging, and even cancer. Understanding the functional evolution of the hydrozoan genes related to reverse development, will provide new insights into the genetic and molecular mechanisms that may potentially be applied to future medical research.


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