Cover. Co-expression network of transcripts of Nostoc sp. PCC 7120 with expression changes after removal of combined nitrogen (left). 3D structural model of sRNAs Yfr1 (top right) and NsiR1 (bottom right). Models were generated by RNAComposer (Popenda et al. 2012. Nucleic Acids Research, 40: e112. doi: 10.1093/nar/gks339).

Instituto de Bioquímica Vegetal y Fotosíntesis

Departamento de Bioquímica Vegetal y Biología Molecular

Universidad de Sevilla - Consejo Superior de Investigaciones Científicas

TESIS DOCTORAL

Global identification of regulatory RNAs in the cyanobacterium Nostoc sp. PCC 7120. Functional characterization of Yfr1 and NsiR1.

Trabajo presentado por Manuel Brenes Álvarez para optar al grado de Doctor

Manuel Brenes Álvarez,

Sevilla, 2019

Director Directora

Dr. Agustín Vioque Peña Dra. Alicia María Muro Pastor

Catedrático de la Universidad de Sevilla Científica Titular del Consejo Superior de Investigaciones Científicas

FUNDING

This Doctoral Thesis has been carried out at the Instituto de Bioquímica Vegetal y Fotosíntesis (IBVF, CSIC-US) and has been funded by a FPU predoctoral contract from Ministry of Education, Culture and Sport (FPU014/05123). The work has been funded by projects from Ministry of Economy and Competitiveness (BFU2013-48282-C2-1-P, Non-coding RNAs involved in adaptation to nitrogen stress and cell differentiation in cyanobacteria) and from State Research Agency, Ministry of Economy, Industry and Competitiveness (BFU2016- 74943-C2-1-P, Participation of non-coding RNAs in regulatory circuits controlled by nitrogen availability in cyanobacteria) whose Principal Investigator was Dr. Alicia María Muro Pastor. Both projects were co-financed by the European Regional Development Fund (ERDF). In addition, a three-months stay at the laboratory of Dr. Wolfgang Hess (University of Freiburg, Germany) was funded by a fellowship from Ministry of Education, Culture and Sport (EST16/00088).

A mis abuelos

AGRADECIMIENTOS

En primer lugar, me gustaría agradecer a mis directores, Agustín y Alicia, su paciencia y dedicación en estos años que hemos trabajado juntos. Su pasión por la Ciencia y su constante atención han sido una inspiración y un importante apoyo a lo largo de esta tesis.

Al Dr. José Enrique Frías, por su experta ayuda durante la realización de los ensayos de actividad de nitrato y nitrito reductasa. A la Dra. Iris Maldener y la Dra. Rebeca Perez por su ayuda en las tinciones de peptidoglicano naciente.

I would like to thank Dr. Wolfgang Hess for allowing me to stay for three months in his laboratory in Freiburg. Thank you for being so kind and helpful, making me feel like home there.

Gracias a toda mi familia por su inmenso cariño y apoyo. A mis abuelos, por enseñarme que con trabajo, constancia y esfuerzo todo se consigue. A mis padres, porque sin su educación e inmensa confianza probablemente esta tesis no habría tenido lugar. A mi hermana Mercedes, porque su valentía para enfrentarse a nuevos problemas siempre ha sido un motivo de inspiración.

A mis compañeros de laboratorio, Isidro y Elvira, por su ayuda y por hacer mucho más ameno todo mi tiempo en el laboratorio. A Javi, Alejandro, Mari, Diego, Tommy y muchos otros compañeros, por no ser sólo compañeros de trabajo, sino también buenos amigos con los que salir, charlar, desconectar del trabajo y, en resumen, disfrutar de la vida.

Por último, quisiera darle las gracias a Ana por apoyarme en todo lo que hago. Gracias por escucharme detenidamente cuando comento mis tribulaciones científicas, incluso aunque el tema te sea completamente desconocido y gracias por enseñarme cómo no debo dejarme abrumar por los problemas.

Finalmente, gracias a todas aquellas personas que de un modo u otro han influido en mi desarrollo como persona y en la consecución de este trabajo de tesis.

We ought not to hesitate nor to be abashed, but boldly to enter upon our researches concerning animals of every sort and kind, knowing that in not one of them is Nature or Beauty lacking.

Aristotle. Parts of animals, I.

INDEX

INDEX

1. INTRODUCTION ...... 1

1.1 The cyanobacteria ...... 3 1.1.1 General characteristics ...... 3 1.1.2 Cyanobacterial cell envelope ...... 7 1.1.2.1 Synthesis and remodeling of peptidoglycan ...... 7 1.1.2.2 Transporters ...... 9 1.1.3 Metabolic characteristics ...... 10 1.1.3.1 Carbon assimilation in cyanobacteria ...... 11 1.1.3.2 Nitrogen assimilation in cyanobacteria ...... 11 1.1.4 Adaptation to combined nitrogen deprivation ...... 15 1.1.4.1 General response ...... 15 1.1.4.2 Heterocyst differentiation...... 16

1.2 RNAs regulators in bacteria ...... 20 1.2.1 Types of RNA regulators ...... 23 1.2.1.1 RNA regulators encoded in cis ...... 23 1.2.1.2 RNA regulators encoded in trans ...... 24 1.2.2 Identification of RNA regulators and their targets ...... 27 1.2.3 RNA regulators in cyanobacteria ...... 28 1.2.3.1 Identification of RNA regulators in cyanobacteria ...... 28 1.2.3.2 Physiological processes regulated by RNA regulators in cyanobacteria ...... 29

2. OBJECTIVES...... 33

3. SUMMARY OF RESULTS ...... 37

3.1 CHAPTER I: A computational approach for the identification of conserved sRNAs in heterocyst-forming cyanobacteria...... 45 3.2 CHAPTER II: Yfr1, a widely conserved sRNA, regulates the integrity of the cell wall and its remodeling during heterocyst differentiation ...... 67 3.3 CHAPTER III: A co-expression network to dissect the complex transcriptome of Nostoc sp. PCC 7120 during heterocyst differentiation...... 103 3.4 CHAPTER IV: NsiR1, a sRNA with multiple copies, regulates heterocyst differentiation . 157

4. GENERAL DISCUSSION ...... 209

5. CONCLUSIONS ...... 221

6. GENERAL REFERENCES ...... 225

i

1. INTRODUCTION

Introduction

1. INTRODUCTION

1.1 The cyanobacteria

1.1.1 General characteristics

Cyanobacteria are Gram-negative bacteria that form a monophyletic group inside the eubacteria (Woese, 1987). They are photosynthetic microorganisms with a photosynthetic apparatus similar to that of chloroplast of algae and higher plants (DeRuyter and Fromme, 2008) and are the only prokaryotes able to perform oxygenic photosynthesis. Nowadays, it is widely accepted that chloroplasts from algae and higher plants have evolved from ancient cyanobacteria that established a symbiotic relationship with a phagotrophic eukaryote (Margulis, 1975; Ochoa de Alda et al., 2014). However, there are some differences between the photosynthetic apparatus of cyanobacteria and algae and those of higher plants. For example, in contrast to algae and higher plants, most cyanobacteria do not have chorophyll b (Stanier and Cohen-Bazire, 1977) and contain phycobilisomes as supramolecular light-harvesting complexes (Grossman et al., 1993b).

Cyanobacteria played a crucial role in Earth’s history. They are thought to be the first organisms that performed an oxygenic photosynthesis (Buick, 1992). The oldest cyanobacterial fossils were found in sedimentary rocks generated more than 3500 million years ago (Schopf and Packer, 1987) and, although with some controversy, it is now accepted that these first photosynthetic organisms may have appeared between 2350 (Kirschvink and Kopp, 2008) and 3600-3700 million years ago (Garcia-Pichel et al., 2019). The ability to use water as electron donor, and the production of oxygen as a result, contributed to one of the most drastic changes in the Biosphere, the change from an anoxygenic atmosphere to an oxygenic atmosphere, a process known as the Great Oxidation Event (GEO) (Lyons et al., 2014). This event allowed the subsequent evolution of heterotrophic aerobic metabolism (Falkowski et al., 2005). In addition, cyanobacteria are also very relevant in the actual Biosphere. Not only is a substantial amount of

CO2 fixation carried out by cyanobacterial photosynthesis in the oceans, but, given that some cyanobacteria are also able to fix atmospheric nitrogen, this group of organisms plays an important role in the carbon, nitrogen and oxygen cycles (Montoya et al., 2004).

Cyanobacteria have colonized diverse ecological niches, from oceans, fresh water and soil to extreme environments such as hot springs, deserts and artic regions. In addition, some species are able to differentiate several cell types or to live in a symbiotic relationship with fungi, plants or protists (Adams, 2002; Meeks and Elhai, 2002). This ability to adapt to environments

3 Introduction

with variable light intensity, temperature or nutrient availability could imply the existence of specific regulatory mechanisms in these bacteria, in addition to more general mechanisms equivalent to those found in other prokaryotes (Tandeau de Marsac and Houmard, 1993).

The wide variety of morphologies shown by cyanobacteria was one of the first criteria used for its classification (Rippka et al., 1979), leading to a classification in five sections (Figure 1).

Figure 1. Representative cyanobacterial strains from the sections in the morphological classification of Rippka et al., 1979. Numbers indicate the taxonomic sections; (I) Chroococcales, (II) Pleurocapsales, (III) Oscillatoriales, (IV) Nostocales and (V) Stigonematales. Images are courtesy of Alicia M. Muro (I, II), José Enrique Frías, Servicio de Cultivos Biológicos del Centro de Investigaciones Científicas Isla de la Cartuja (cicCartuja) (III, V) and Elvira Olmedo (IV).

Section I (Chroococcales), is formed by unicellular cyanobacteria that reproduce by binary fission or by budding (e.g. genera Synechocystis or Synechococcus). Section II (Pleurocapsales), including genus Pleurocapsa, is formed by unicellular cyanobacteria that reproduce by multiple fission. Section III (Oscillatoriales) is formed by filamentous strains that do not differentiate specialized cells (e.g. genera Oscillatoria or Spirulina). Section IV (Nostocales), including genera Nostoc and Anabaena, and Section V (Stigonematales), including genus Fischerella, are formed by filamentous strains that have the capacity of cellular differentiation. Strains with cell division taking place in one plane, therefore unbranched, are classified in Section IV whereas branched strains, in which cell division may take place in more than one plane, are classified in Section V. Strains in these two last sections are able to differentiate several cell types. They can differentiate heterocyst (specialized cells that fix atmospheric nitrogen) under nitrogen deprivation or akinetes (a spore-like cell) under certain stress conditions. In addition, some strains can differentiate hormogonia, small motile filaments

4 Introduction that participate in the spreading of the organism and the establishment of symbiotic relationships (Flores and Herrero, 2010).

More recent classifications are based on methods of molecular phylogeny. These methods have shown that the morphological classification is not coherent with the phylogeny (Giovannoni et al., 1988; Turner et al., 1999; Sánchez-Baracaldo et al., 2005; Tomitani et al., 2006) and only sections IV and V, that include the strains with the capacity of cellular differentiation, are phylogenetically coherent (Sánchez-Baracaldo et al., 2005; Tomitani et al., 2006; Schirrmeister et al., 2015). In 2016, when our approach for the prediction of sRNAs was carried out, the genomes available were basically those included in the analysis of Shih et al., 2013. Therefore, we have used the phylogenetic tree in Shih et al., 2013 as reference (Figure 2). Some conclusions derived from inspection of this phylogenetic tree are a) multicellularity has emerged several times along the evolution of cyanobacteria, and b) some unicellular species seem to be the result of a loss of multicellularity (Shih et al., 2013).

Cyanobacterial genomes show diverse sizes, organization and ploidy. Genes conserved in all cyanobacterial genomes are known as core genes. These genes are related to important and essential physiological process, such as translation, photosynthesis, CO2 fixation and synthesis of chlorophyll and ATP. Along the evolution of cyanobacteria, strains with more complex physiology have acquired specific genes by duplication and genetic drift or by horizontal transfer (Shi and Falkowski, 2008). As a consequence of the acquisition and duplication of genes along the evolution of cyanobacteria, the size of cyanobacterial genomes varies from 1.66 Mb in Prochlorococcus MED4, considered the minimal size of the genome for a non-symbiotic photosynthetic organism (Rocap et al., 2003), to 9.2 Mb in Nostoc punctiforme. The GC content also varies widely between different strains, from 32% to 71%, similar to the GC content range found in prokaryotes as a whole (Herdman et al., 1979). Some cyanobacteria, like Prochlorococcus or Synechococcus, have only one circular chromosome. Other cyanobacteria, like our model organism Nostoc sp. PCC 7120, have their genomic content arranged in a circular chromosome and several plasmids, and some strains even have linear chromosomes, as Cyanothece 51142 (Hess, 2011). Finally, polyploidy is another source of genome heterogeneity. There are monoploid, oligoploid and polyploid strains with more than 100 copies of their chromosome, as some strains of Synechocystis (Griese et al., 2011).

5 Introduction

Figure 2. Cyanobacterial phylogenetic tree according to Shih et al., 2013. Branches are coloured according to the sections in the classification of Rippka et al., 1979 (colour code top right). Framed amplified area contains heterocyst- forming strains, including branched (purple bar) or non-branched strains (pink bar). Nostoc sp. PCC 7120, the model organism used in this work, is indicated by a red arrow. The image, adapted from Shih et al., 2013, shows cyanobacterial genomes sequenced in that work (red). Morphological transitions are indicated by blue triangles.

6 Introduction

The cyanobacterium used as model organism in this Thesis is Nostoc sp. PCC 7120 (a.k.a Anabaena sp. PCC 7120), a non-branched filamentous cyanobacteria with the capacity to differentiate heterocysts (Section IV). The genome of Nostoc sp. PCC 7120 is distributed in a circular chromosome of 6,4 Mb and six plasmids with a size between 405 kb and 5.5 kb, named α, β, γ, δ, ε and ζ (Kaneko et al., 2001).

1.1.2 Cyanobacterial cell envelope

The cell wall of cyanobacteria resembles a Gram-negative architecture, with a peptidoglycan layer between an inner cytoplasmic membrane and an outer membrane (Silhavy et al., 2010). However, cyanobacterial cell envelopes have some unusual features. The peptidoglycan layer is relatively thick and has a high-degree of crosslinking, typical features of Gram-positive strains (Hoiczyk and Hansel, 2000). In addition, the outer membrane in multicellular cyanobacteria does not enter the septa between adjacent cells (Wilk et al., 2011), generating, as a consequence, a continuous periplasm (Mariscal et al., 2007).

1.1.2.1 Synthesis and remodelling of peptidoglycan

Peptidoglycan is a strong and flexible mesh that surrounds the cell, contributes to its shape and protects it against osmotic pressure. Peptidoglycan contains glycan strands of alternating N-acetylglucosamine (GluNAc) and N-acetylmuramic acid (MurNAc) cross-linked by short peptides (Vollmer et al., 2008a). A pentapeptide substitutes the D-lactoyl group of each MurNAc residue and, in Gram-negative strains, usually has the sequence L-Ala-γ-D-Glu-meso- 2,6-diaminopimelic-acid-D-Ala-D-Ala. The last D-Ala residue is lost when the D-Ala at position 4 of the pentapeptide in one glycan strand is crosslinked to the diaminopimelic acid at position 3 of the pentapeptide in another glycan strand.

The biosynthesis of peptidoglycan involves three main steps; the synthesis of the disaccharide with the pentapeptide in the cytosol, the translocation of this disaccharide- pentapeptide unit across the plasma membrane and the transglycosylation and transpeptidation of new units in the nascent peptidoglycan strands (Figure 3). The first step of peptidoglycan synthesis is the conversion of GluNAc to MurNAc. Later, the sequential addition of amino acids (including D-Ala generated by alanine racemase) to the nascent pentapeptide

7 Introduction

ligated to MurNAc is operated by different Mur ligases. The completed MurNAc-pentapeptide is then ligated by MraY to a membrane-bound undecaprenyl-phosphate carrier to produce lipid I, and later GluNAc is added to produce lipid II. The lipid II-bound disaccharide with the pentapeptide is translocated from the inner side of the plasma membrane to the periplasmic space where it is added to the nascent peptidoglycan by the transglycosylase and transpeptidase activities of several penicillin-binding proteins (PBP).

Figure 3. Peptidoglycan biosynthesis. The biosynthetic pathway has three main steps, the cytosolic synthesis of the disaccharide with the pentapeptide, the lipid-linked steps that allow the translocation of the subunit across the membrane and the incorporation of the new subunit into the peptidoglycan mesh by transglycosylases and transpeptidases. Taken from Nachar et al., 2012.

The sacculus of peptidoglycan is a dynamic structure that must adapt to different physiological processes, such as the turnover of peptidoglycan, the assembly of large trans- envelope secretion systems, the growth of the cells or the separation of daughter cells during cell division. This constant remodeling is performed by different murein hydrolases, such as N- Acetylmuramoyl-L-Ala amidases (Ami enzymes), reviewed in Vollmer et al., 2008b.

The profound morphological changes that heterocysts undergo during the differentiation process (see section 1.1.4.2) involve also changes in the sacculus of peptidoglycan. Mutants in peptidoglycan biosynthesis enzymes, as MurC (alr5065) and MurB (alr5066) (Videau et al., 2016b), or in penicillin binding proteins (all2981, alr4579 and alr5051) (Lázaro et al., 2001; Leganés et al., 2005) show alterations in the differentiation of heterocyst

8 Introduction and growth defects in media without combined nitrogen. In addition, the growth of a filament in media without combined nitrogen relies in the exchange of nitrogen and carbon compounds between vegetative cells and heterocysts (see 1.1.4.2). These metabolites may diffuse along the filament from cytoplasm to cytoplasm through septal junctions (Mullineaux et al., 2008; Nürnberg et al., 2015; Nieves-Morión et al., 2017; Weiss et al., 2019), proteinaceous complexes that traverse the septal peptidoglycan disks through perforations (nanopores) made by murein amidases (Nürnberg et al., 2015). Mutants of murein amidases AmiC1 (alr0092) (Bornikoel et al., 2018) or AmiC2 (alr0093) (Zhu et al., 2001; Bornikoel et al., 2017) show less nanopores and defects in diazotrophic growth.

1.1.2.2 Transporters

The cytoplasmic and outer membranes are selective barriers that prevent the free diffusion of substances to the cytosol (Nicolaisen et al., 2009b). Different classes of transporters facilitate movement of metabolites through these two membranes. Some transporters are located in the outer membrane, others are located in the cytoplasmic membrane and some secretion systems are large complexes that form channels that allow the direct secretion of molecules from the cytosol to the exterior (Nicolaisen et al., 2009a).

General porins, OprB-like porins and TonB-dependent transporters are located in the outer membrane. General porins are large channels, usually with a low selectivity, that allow the diffusion of hydrophilic molecules of a size up to 600 Da (Koebnik et al., 2000). However, OprB- like and TonB-dependent transporters have some selectivity; OprB-like porins may be involved in the uptake of carbohydrates (Nicolaisen et al., 2009a) and TonB-dependent transporters are specifically involved in the uptake of siderophores (Noinaj et al., 2010).

ABC transporters are located in the cytoplasmic membrane and transport diverse molecules using ATP as energy source (Shvarev and Maldener, 2018). Some transporters, such as transporters for nitrate, nitrite or urea, are regulated by nitrogen availability (see section 1.1.3.2), and others are specifically involved in the secretion of components of heterocyst envelopes, as HepA (Holland and Wolk, 1990; Zhu et al., 1998), DevBCA (Maldener et al., 1994; Fiedler et al., 1998) or HgdBC (Shvarev et al., 2018).

Finally, a putative type 1 secretion system formed by DevBCA and HgdD has been reported (Staron et al., 2011). This system could directly export heterocyst glycolipids (HGL, see

9 Introduction

1.1.4.2) from the cytosol to the heterocyst envelope outside the outer membrane thanks to the presence of the TolC-like protein HgdD. This protein could form a promiscuous channel that, through interaction with different transport systems, may participate in the efflux of different substances (Hahn et al., 2012; Hahn et al., 2015).

1.1.3 Metabolic characteristics

In contrast to their diverse morphology and ecological distribution, most cyanobacteria share a similar central metabolism. Cyanobacteria perform oxygenic photosynthesis, although some species, like Oscillatoria limnetica, under certain conditions, are

able to perform an anoxygenic photosynthesis using SH2 as electron acceptor generating S (Shahak et al., 1987). Cyanobacteria were the first organisms to evolve two photosystems working in series. This innovation generated an electrochemical potential that allowed ancient cyanobacteria to use water as electron acceptor and, consequently, generate oxygen as a product of the reaction (Hohmann-Marriott and Blankenship, 2011). Oxygenic photosynthesis is restricted to cyanobacteria, algae and higher plants and the process is essentially the same in these organisms. However, as it was mentioned before, cyanobacteria have some peculiarities. Cyanobacteria contain phycobilisomes, supramolecular light harvesting complexes formed by phycobiliproteins and antenna pigments that channel and transfer light energy to the photosystems (Grossman et al., 1993b). In addition, these organisms are able to modify their photosynthetic apparatus in response to different light conditions, in a process called chromatic adaptation. For example, they can change the content of pigments in phycobilisomes in response to changes in wavelength or light intensity (Gutu and Kehoe, 2012; Montgomery, 2016).

In cyanobacteria, photosynthesis and respiration coexist in the same compartment, therefore some components of the respiratory and photosynthetic chains are shared by both processes (Schmetterer, 2006). For many years, the cyanobacterial Krebs cycle was considered incomplete because they lack 2-oxoglutarate dehydrogenase (Stanier and Cohen-Bazire, 1977). This incomplete cycle could have an anabolic function in the incorporation of inorganic nitrogen to carbon chains or could participate in the biosynthesis of amino acids (as glutamate), porphyrins, phycobilins and chlorophyll (Beale, 1994). However, recent studies have suggested the existence of alternative pathways that could close the Krebs cycle (Zhang and Bryant, 2011; Xiong et al., 2014).

10 Introduction

1.1.3.1 Carbon assimilation in cyanobacteria

Cyanobacteria were classically considered strict autotrophs that only use CO2 as carbon source. However, due to some metabolic capacities or the existence of transporters and permeases that facilitate the uptake of sugars (Ungerer et al., 2008), some cyanobacteria are able to use reduced carbon compounds as a source of carbon (Rippka, 1972). Nowadays, several examples of cyanobacteria with a photochemoheterotrophic, chemoheterotrophic or mixotrophic metabolism have been reported (Pelroy et al., 1972; Stebegg et al., 2012; Muñoz- Marín et al., 2013).

Fixation of CO2 is carried out by RuBisCO (ribulose-1,5-bisphosphate carboxylase/oxygenase), considered the most abundant enzyme on Earth. RuBisCO is a bifunctional enzyme that can use CO2 (carboxylase activity) or O2 (oxygenase activity).

Carboxylase activity leads to incorporation of one molecule of CO2 to one molecule of ribulose 1,5-bisphosphate generating two molecules of 3-phosphoglycerate. This metabolite enters the biosynthetic metabolism by the Calvin-Benson cycle and part of the fixed CO2 can be accumulated as glycogen, as a reduced carbon storage form for periods of darkness. However, when RuBisCO acts as oxygenase, ribulose 1,5-bisphosphate is oxidized generating 2- phosphoglycolate, a metabolite that requires great amount of energy to be metabolized by a process known as photorespiration. Given that the concentration of O2 in the atmosphere is greater than the concentration of CO2, and O2 is generated during photosynthesis, photorespiration is inevitable and decreases the efficiency of photosynthesis (Kroth, 2015). In order to increase the local concentration of CO2 around the RuBisCO, cyanobacteria use several strategies (Burnap et al., 2015), including a bacterial microcompartment called carboxysome that encapsulates carbonic anhydrase and RuBisCO, favouring fixation of CO2 (Kerfeld and Melnicki, 2016).

1.1.3.2 Nitrogen assimilation in cyanobacteria

Nitrogen is one of the most abundant bioelements. Cyanobacteria are able to use different sources of nitrogen, such as ammonium, nitrate, nitrite, nitrogenous bases and some amino acids as glutamine and arginine (Flores and Herrero, 1994). However, similar to the catabolite repression exerted in Escherichia coli, cyanobacteria use preferentially those nitrogen

11 Introduction

sources whose assimilation requires less energy, a process called “nitrogen control” (Herrero et al., 2001; Herrero and Flores, 2019).

Ammonium is the preferred nitrogen source, so that when ammonium is present the genes involved in the assimilation of alternative nitrogen sources are repressed (Herrero and Flores, 2019). Ammonium is directly incorporated in carbon skeletons whereas the rest of nitrogen sources need to be intracellularly converted to ammonium before being incorporated to the biosynthetic pathways (Figure 4). Amt1, a member of the Amt family of widely distributed transporters acting as ammonium scavengers (Wacker et al., 2014), seems to be the main ammonium transporter in cyanobacteria (Montesinos et al., 1998).

Figure 4. Nitrogen assimilation in undifferentiated cells of Nostoc sp. PCC 7120. Ammonium translocator (Amt) and ABC transporters for nitrate and nitrite (Nrt) or urea (Urt) mediate uptake of nitrogenous compounds. The nitrate/nitrite assimilation pathway involves two sequential reductions by the reduced ferredoxin (Fdred)-dependent enzymes nitrate reductase (NarB) and nitrite reductase (NirA). Urea is reduced to ammonium by the multienzymatic urease complex (UreABC). Finally, ammonium is incorporated to glutamate by glutamine synthetase (GS), generating glutamine, whose amide group is later transferred to one molecule of 2-oxoglutarate (2-OG) by glutamate synthase (GOGAT), producing two molecules of glutamate. Adapted from Herrero and Flores, 2019.

Ammonium is incorporated by the GS-GOGAT cycle (Wolk et al., 1976; Flores and Herrero, 1994; Meeks and Elhai, 2002), which is the link between the carbon and nitrogen cycles (Zhang et al., 2018). The first step of the GS-GOGAT cycle is the synthesis of glutamine by glutamine synthetase (GS) using ammonium and glutamate, a step that requires energy in the form of ATP. The second step is catalysed by the enzyme glutamate synthase (GOGAT) that transfers the amide group of glutamine to a molecule of 2-oxoglutarate, generating two molecules of glutamate in a reaction that needs reductants (van den Heuvel et al., 2004). Finally, glutamate is used for the recycling of glutamine, necessary for the continuity of the cycle.

12 Introduction

The global regulator of nitrogen assimilation in cyanobacteria is NtcA, a transcriptional regulator of the CRP/FNR family that can function as an activator or repressor of the transcription of genes regulated by nitrogen availability (Herrero et al., 2001; Luque and

Forchhamer, 2008). NtcA binds to a specific sequence, first defined as GTAN8TAC based on a few promoters (Luque et al., 1994), and later redefined as GTN10AC based in global approaches that consider larger sets of promoters regulated by this transcription factor (Mitschke et al., 2011b; Picossi et al., 2014; Giner-Lamia et al., 2017). In most activated promoters, the NtcA binding site is centred around position -41.5 with respect to the transcription start site (TSS), similar to the position of CRP binding sites in the Class II activated promoters, allowing the productive interaction with RNA polymerase (Herrero et al., 2001; Browning and Busby, 2004; Mitschke et al., 2011b). However, in repressed promoters, the NtcA binding site overlaps the -35 box, the - 10 box or the TSS region, therefore interfering the binding of the RNA polymerase to the promoter (Herrero et al., 2001). In addition, the affinity of NtcA to its binding site, and consequently its activity, is modulated by the concentration of 2-oxoglutarate, an indicator of the nitrogen status of the cell (Muro-Pastor et al., 2001). Under nitrogen deprivation, there is an increase in the concentration of 2-oxoglutarate, a molecule that binds to NtcA in a pocket structurally similar to the one involved in the binding of cAMP to CRP (Zhao et al., 2010) and promotes the binding of NtcA to its binding sites (Valladares et al., 2008).

NtcA activity is also modulated by proteins PII and PipX (PII interacting protein X). PII is a protein widely conserved in bacteria, algae and plants and senses changes in the availability of carbon, energy or nitrogen (Forcada-Nadal et al., 2018). This protein was first described in E. coli (Adler et al., 1975) as a 2-oxoglutarate sensor that regulates several process through protein- protein interactions. In cyanobacteria, PII modulates the biosynthesis of arginine, the transport of nitrate and nitrite and expression of genes regulated by NtcA (Luque and Forchhamer, 2008).

PipX is only conserved in cyanobacteria and interacts with PII but also with NtcA, depending on 2-oxoglutarate availability (Espinosa et al., 2006). PipX acts as a coactivator of NtcA (Espinosa et al., 2006; Zhang et al., 2018). Under combined nitrogen deprivation, the high concentration of 2-oxoglutarate stabilizes the PipX-NtcA complex, favouring the activity of NtcA, and provokes a conformational change in PII that prevents sequestration of PipX by PII. So, the complex network of interactions between NtcA, PII and PipX modulates the response to nitrogen availability (Forcada-Nadal et al., 2018). NtcA is also essential for the differentiation of heterocysts (see section 1.1.4.2) and PipX is needed for proper function of heterocysts. PipX is needed for the full induction of nitrogenase genes and genes related to the protection against oxygen (Valladares et al., 2011). The induction of pipX (asr0485) in Nostoc sp. PCC 7120 under nitrogen deprivation

13 Introduction

depends on NtcA but also on HetR, the master regulator of heterocyst differentiation (see section 1.1.4.2).

Cyanobacteria can also use organic compounds as nitrogen source. For example, Nostoc sp. PCC 7120 can use glutamine, arginine (Burnat and Flores, 2014; Burnat et al., 2018) or urea (Valladares et al., 2002). However, nitrogen sources are used hierarchically, with inorganic nitrogen sources, as nitrate or nitrite, used preferentially. Nitrate or nitrite enter the cells by high affinity transporters and are reduced in two successive reactions that use reduced ferredoxin as electron donor, linking nitrogen assimilation to photosynthesis (Flores et al., 2005). Nitrate is reduced to nitrite, nitrite is reduced to ammonium and ammonium is finally assimilated by the GS-GOGAT cycle (Figure 4). Gene products involved in nitrate assimilation are encoded in the nir operon, whose transcription is activated by NtcA (Figure 5) (Frías et al., 1997). This operon encodes nitrite reductase (NirA), the subunits of one of the main nitrate/nitrite transporters (NrtABCD) and nitrate reductase (NarB). In addition, NtcA also activates the expression of another operon that encodes NtcB and CnaT, two proteins necessary for the full induction of the nir operon. NtcB is a transcription factor of the LysR family that binds to the promoter region and promotes the transcription of the nir operon (Frías et al., 2000) and CnaT boosts the expression of the nir operon by an unknown mechanism (Frías et al., 2003). The activity of nitrate assimilation enzymes is also regulated by other factors. For example, proteins NirB and NarM seem to interact with NirA and NarB respectively (Frías and Flores, 2015), and these interactions are necessary for full activity of these enzymes (Frías and Flores, 2010; 2015). Finally, the NirA/NirB complex seems to negatively affect the nir operon expression when nitrate/nitrite are not present in the media (Frías and Flores, 2010).

Figure 5. Regulation of the nitrate assimilation cluster. NtcA bound to 2-oxoglutarate (2-OG) activates the expression of the nir operon encoding nitrite reductase (nirA), nitrate reductase (narB) and nitrate/nitrite transporter (nrtABCD). ntcB and cnaT are also induced by NtcA+2-OG and their products are necessary for the full activation of the nir operon; NtcB binds to the promoter of the nir operon and activates its expression whereas CnaT enhances the expression of the nir operon by an unknown mechanism. NirB and NarM are also necessary for the proper activity of nitrite reductase (NirA) and nitrate reductase (NarB), respectively. Finally, in the absence of nitrate or nitrite, the complex NirA/NirB seems to negatively affect the expression of the nir operon. Taken from Herrero and Flores, 2019.

14 Introduction

1.1.4 Adaptation to combined nitrogen deprivation

1.1.4.1 General response

When cyanobacteria sense a lack of combined nitrogen, a general response takes place to try to overcome such lack of nutrients. This response is under control of NtcA and common to all cyanobacteria, whether or not they are ultimately able to fix atmospheric nitrogen, a process that in some strains requires differentiation of heterocysts (see below). This general response involves induction of the expression of transporters of ammonium (Flaherty et al., 2011; Mitschke et al., 2011b) nitrate/nitrite (Frías et al., 1997) and urea (Valladares et al., 2002). Expression of the genes required for the assimilation of these alternative sources is also induced, in order to use traces of nitrate, nitrite, urea, amino acids or even ammonium that may remain in the media. In addition, the activity of glutamine synthetase (GS, encoded by glnA) is induced by several mechanisms. NtcA promotes the transcription of glnA (Herrero et al., 2001), inhibits the transcription of the inactivating factors of glutamine synthetase IF7 and IF17 (encoded by gifA and gifB) (García-Domínguez et al., 2000) and promotes the transcription of NsiR4, a sRNA that inhibits translation of IF7 (Klähn et al., 2015). In addition, a glutamine riboswitch regulates the expression of the other inactivating factor (IF17) (Klähn et al., 2018).

In the absence of combined nitrogen, cyanobacteria activate the degradation of proteins (Fleming and Haselkorn, 1974). Therefore, another aspect of the general response to nitrogen deprivation is the recycling of the amino acids present in the proteins that constitute the phycobilisomes, that represent more than 40% of the cellular protein and constitute a major storage of carbon and nitrogen. Under nitrogen deprivation, these supramolecular complexes are dismantled, in a process known as “bleaching” (Grossman et al., 1993b; 1993a), and their amino acids are recycled and incorporated in other metabolic pathways. NblA (non-bleaching phenotype A) is a small protein that plays an important role in this process mediating the interaction between the phycobiliproteins and the proteases that degrade them (Collier and Grossman, 1994; Karradt et al., 2008; Sendersky et al., 2014). Phycobilisome degradation is a general response common to all cyanobacteria but differs in strains that are eventually able to fix atmospheric nitrogen from those that are not. Whereas in non-N2-fixing cyanobacteria, phycobilisome degradation persists in time, in N2-fixing cyanobacteria, such as our model organism, phycobilisome degradation is only a transient response until heterocysts start to fix

N2 (Baier et al., 2004).

15 Introduction

1.1.4.2 Heterocyst differentiation

Some cyanobacteria have the capacity of fix atmospheric nitrogen by nitrogenase, a

multiproteic complex that reduces N2 to ammonium in a reaction that consumes a high amount of energy (Seefeldt et al., 2009). Because nitrogenase is extremely sensitive to oxygen, cyanobacteria have developed several strategies to solve the incompatibility between nitrogen fixation and oxygenic photosynthesis (Fay, 1992). Non heterocyst-forming cyanobacteria usually alternate these processes, fixing nitrogen during darkness periods and performing oxygenic photosynthesis during daytime (Huang et al., 1999; Reade et al., 1999; Misra and Tuli, 2000). Heterocyst-forming cyanobacteria can spatially separate these two processes. The differentiation of some vegetative cells into heterocysts in a semi-regular pattern generates a multicellular organism with two specialized cell types, vegetative cells and heterocysts.

Vegetative cells perform photosynthesis and CO2 fixation whereas heterocysts provide a micro-

oxic environment for N2 fixation. So, both cell types need to exchange carbon and fixed nitrogen in order to sustain the growth of the filament as whole (Figure 6) (Muro-Pastor and Hess, 2012).

Figure 6. Heterocyst differentiation and metabolic exchange between vegetative cells and heterocysts. (A) Filaments of Nostoc sp. PCC 7120 growing on top of plates without combined nitrogen differentiate new heterocysts (white triangles) between two mature heterocysts (black triangles). (B) Approximate timing of heterocyst differentiation. (C) Division of labour and metabolic exchange between heterocysts (nitrogen fixation) and vegetative cells (oxygenic photosynthesis and carbon fixation). Taken from Muro-Pastor and Hess, 2012.

16 Introduction

Heterocysts are terminally differentiated non-dividing cells devoted to nitrogen fixation. Its morphology and physiology have evolved to generate a micro-oxic environment that protects and supports nitrogenase activity. Heterocyst differentiation involves a complex developmental program that includes morphological changes, metabolic reprogramming, regulation of pattern and timing of differentiation and commitment (Flores et al., 2019).

Heterocyst are morphologically different from vegetative cells. They are larger and have a thick envelope that restricts gas diffusion and constraints the contact area between heterocysts and vegetative cells (Fay, 1992). Their thylakoid membranes are reorganized at the poles in a structure called “honeycomb” (Lang and Fay, 1971), they don’t have carboxysomes (Winkenbach and Wolk, 1973; Stewart and Codd, 1975) and they accumulate fixed nitrogen in granules of cyanophycin localized at the poles (Sherman et al., 2000).

The micro-oxic environment of heterocysts is achieved through several strategies. Heterocyst envelopes contain two extra-layers outside the cell wall; an outer polysaccharide layer (HEP) (Bryce et al., 1972) and an inner glycolipid layer (HGL) (Cardemil and Wolk, 1979). Genes required for the synthesis of the HEP and HGL layers are mainly encoded in clusters called “gene expression islands”. Most of the genes related to synthesis of the HEP layer are encoded in one expression island (alr2825-alr2841) (Huang et al., 2005), but some are also outside of the cluster (Maldener et al., 2003; Wang et al., 2007), and are transcribed at early stages in the heterocyst differentiation process (Ehira et al., 2003; Flaherty et al., 2011; Mitschke et al., 2011b). Most of the genes involved in the biosynthesis and deposition of the components of the HGL layer are clustered in another “gene expression island” (all5341- all5359) (Fan et al., 2005; Awai and Wolk, 2007) and are transcribed later during the differentiation process (Ehira et al., 2003; Flaherty et al., 2011). Thus, the HEP layer is deposited first, and at an early stage, and the HGL layer is deposited beneath it. In addition to the specialized heterocyst envelope, several enzymatic activities contribute to reducing the intracellular concentration of oxygen. Heterocysts have a different organization of their thylakoid membranes. Vegetative cells usually have their thylakoid membranes concentrically arranged in the periphery whereas, in heterocysts, membranes are localized near the poles (Wildon and Mercer, 1963; Lang and Fay, 1971). Terminal respiratory oxidases localized in these membranes (Cox oxidases) (Valladares et al., 2003) and cytoplasmic flavodiiron proteins (Flv1B and Flv3B) (Ermakova et al., 2014) consume O2 in the heterocyst, contributing to its micro-oxic ambient. In addition, a specific peroxidase that is induced in heterocysts (RbrA) protects these cells from the Reactive Oxygen Species (ROS) (Zhao et al., 2007). Finally, photosystem II (PSII) is not functional in heterocysts

(Donze et al., 1972) and there is no photosynthetic CO2 fixation because of lack of RuBisCO

17 Introduction

(Winkenbach and Wolk, 1973; Stewart and Codd, 1975). However, given that nitrogenase requires a high amount of energy and reductants, heterocysts maintain photosystem I (PSI) and perform photophosphorylation (Magnuson and Cardona, 2016).

Nitrogenase is a multiproteic complex. Genes encoding factors involved in the maturation of nitrogenase, its enzymatic activity or its specific electron donors are encoded in a large cluster (nifB-fdxN-nifS-nifU-nifH-nifD-nifK-nifE-nifN-nifX-all1435-asl1434-nifW-hesA-hesB- fdxH) that is transcribed in the last steps of heterocyst differentiation (Ehira et al., 2003; Flaherty

et al., 2014). Nitrogenase reduces N2 to ammonium that is later assimilated by glutamine synthetase, producing glutamine (Wolk et al., 1976). Because heterocysts don’t have GOGAT activity (Thomas et al., 1977; Martín-Figueroa et al., 2000), glutamine must be transferred to vegetative cells in exchange for glutamate in order to close the GS-GOGAT cycle. So, heterocysts feed vegetative cells with glutamine and other nitrogen compounds (Burnat et al., 2014) and in return, vegetative cells feed heterocysts with reduced carbon compounds. For example, vegetative cells feed heterocysts with sucrose that is hydrolysed in the heterocysts by invertase (InvB) (López-Igual et al., 2010; Vargas et al., 2011), generating fructose and glucose that can be later oxidised by the oxidative pentose-phosphate cycle generating energy and reductant (Magnuson and Cardona, 2016). This exchange of metabolites could not be possible without the structures and pathways of exchange commented in section 1.1.2.

The regulatory cascade that orchestrates heterocyst differentiation is complex and not fully understood. The two essential transcription factors involved in differentiation of heterocysts are NtcA and HetR. Whereas NtcA is the global regulator of nitrogen assimilation (Herrero et al., 2001; Herrero et al., 2004), HetR is specifically involved in cellular differentiation (Buikema and Haselkorn, 1991; Black et al., 1993). At the initiation of heterocyst differentiation, induction of the expression of ntcA and hetR depends on each other (Muro-Pastor et al., 2002) and leads to an overproduction of both factors in cells fated to become heterocysts (Black et al., 1993; Olmedo-Verd et al., 2006). This specific accumulation of HetR and NtcA in some cells may trigger the regulatory cascade that activates the differentiation program. The NtcA-dependent induction of hetR may be mediated by NrrA, a response regulator-like factor whose expression is induced by NtcA (Muro-Pastor et al., 2006) that binds in vitro to the promoter region of hetR (Ehira and Ohmori, 2006).

The expression of hetR is regulated a several levels. hetR is transcribed from a complex promoter that includes constitutive TSS but also one TSS specifically induced in cells becoming heterocysts (Rajagopalan and Callahan, 2010). HetR is a DNA-binding protein (Kim et al., 2011;

18 Introduction

Kim et al., 2013) for which a binding in vitro to its own promoter region and those of several genes specifically induced in heterocyst has been reported (Huang et al., 2004). HetR positively regulates its own expression and the expression of its inhibitors, patS (Yoon and Golden, 1998) and hetN (Callahan and Buikema, 2001), both essential for the maintenance of the pattern of heterocysts along the filaments (Borthakur et al., 2005; Rivers et al., 2018). PatS and HetN contain a RGSGR pentapeptide that binds to HetR, blocks its DNA-binding activity (Huang et al., 2004) and signals its degradation (Risser and Callahan, 2009). The diffusion of PatS (Corrales- Guerrero et al., 2013) or a HetN-derived product may generate a gradient of morphogen that regulates the expression of hetR in the semi-regular pattern that intercalates heterocyst and vegetative cells. HetR is also an auto-proteolytic protein (Zhou et al., 1998) and is degraded by a protease (HetF) (Wong and Meeks, 2001; Risser and Callahan, 2008). In addition, HetR seems to interact with HetP and HetP homologs, proteins that regulate the establishment of the commitment point of heterocysts (Videau et al., 2016a). After this point, pro-heterocysts become irreversibly committed to differentiate while their division machinery gets down- regulated at transcriptional and translational levels (Kuhn et al., 2000; Wang and Xu, 2005).

NtcA and HetR also regulate the early expression of devH, a transcriptional activator of the CRP family that regulates the expression of hgl genes (Hebbar and Curtis, 2000; Ramírez et al., 2005). At an advanced state of maturation, the transcriptional regulator CnfR (previously known as PatB) is specifically induced in heterocysts (Jones et al., 2003). CnfR may be activated by the low oxygen tension inside of heterocyst (Liang et al., 1993), it binds to the promoter region of the nif operon and promotes the expression of nitrogenase genes (Pratte and Thiel, 2016).

In addition, the regulation of heterocyst differentiation involves many other mechanisms, including the use of alternative sigma factors that are specifically expressed in heterocysts at an early (sigC) or late (sigE) stage (Aldea et al., 2007; Muro-Pastor et al., 2017). Finally, several sRNAs that are specifically expressed in heterocysts have been identified (Ionescu et al., 2010; Mitschke et al., 2011b), suggesting a regulatory role of this type of molecules in the complex regulatory network leading to heterocyst differentiation.

19 Introduction

1.2 RNAs regulators in bacteria

The central dogma of molecular biology postulated that the transfer of information in a biological system was unidirectional and finished with the synthesis of proteins (Crick, 1970). In this scenario, the RNA was considered a mere messenger in the transmission of information from the DNA to the synthesis of proteins. However, the use of next generation sequencing techniques has revealed an unexpected amount of non-coding transcripts, apart from ribosomal RNAs (rRNAs) or transfer RNAs (tRNAs). Although the function of some of these non-coding RNAs remains unknown, many of them regulate gene expression (Wade and Grainger, 2014). So, the ubiquity of these molecules and the wide variety of processes that they may regulate suggest an important role for these molecules in the mechanisms of genetic regulation. The mechanisms for the biogenesis of RNA regulators are diverse (Figure 7).

Figure 7. Biogenesis of RNA regulators. (A-C) Antisense RNAs (asRNAs) may be complementary to the coding region, 5’UTR or 3’UTR of their target gene (A). Two mRNAs encoded in opposite orientation may regulate the expression of each other, a type of regulation known as “excludons” (B-C). (D-F) sRNAs may be generated in intergenic regions as independent genes (D), by processing of a mRNA by RNases (orange) (E) or from the 5’UTR, 3’UTR or even inside the coding region of a gene (F). DNA strands are indicated by parallel black lines. TSS are indicated by vertical arrows. Shine Dalgarno sequences (SD) are indicated by green squares.

20 Introduction

One of the main groups of non-coding regulatory RNAs are antisense RNAs (asRNAs) (Lasa et al., 2012), that are transcribed in antisense orientation to the molecule that they regulate (Figure 7A-C). The other main group of non-coding regulatory RNAs are small RNAs (sRNAs), molecules with sizes between 50 and 350 nucleotides that are commonly transcribed from a different genomic region than the molecule(s) that they regulate (Storz et al., 2011). sRNAs may be transcribed from independent genes in intergenic regions (Figure 7D) or they could be generated by the processing of another RNA molecule (Figure 7E), as SdsN (Hao et al., 2016). In addition, they could be transcribed from TSS that are located in the 3’ UTR (UTR, untranslated region) of a gene, as DapZ (Chao et al., 2012), in the coding region of a gene, as MicL (Guo et al., 2014), or they could also share a promoter with a gene, as actuatons (Figure 7F) (Kopf et al., 2015a).

RNA regulators usually exert a post-transcriptional regulation that complements the regulation carried out by transcription factors, playing a role in the fine-tuning necessary for the adaptation of living organism to an ever-changing environment (Wagner and Romby, 2015). The advantages of post-transcriptional regulation exerted by sRNAs versus classical regulation exerted by transcription factors have been widely reviewed (Figure 8) (Beisel and Storz, 2010; Mandin and Guillier, 2013; Wagner and Romby, 2015; Nitzan et al., 2017).

Figure 8. Kinetic properties of sRNA regulation. (A) The regulation exerted by sRNA is faster than the regulation exerted by transcription factors. (B) The 1:1 interaction that occurs in many sRNA-mRNA interactions provokes an inverse relation between free sRNA and free mRNA target. (C) The inverse relation between free sRNA and mRNA target generates a threshold-linear response that filters weak or transient signals. The effective expression of a target does not start until it overpasses the transcription rate of the sRNA (*). After this point, the effective transcription rate of the target depends on the difference between the transcription rate of sRNA and target mRNA. Adapted from Nitzan et al., 2017.

First, because sRNA expression only involves transcription whereas the expression of transcription factors involves transcription and translation, sRNA regulation may be faster than regulation exerted by transcription factors (Figure 8A). In addition, in the case of negative regulation, the binding of a transcription factor to the DNA prevents the transcription of new

21 Introduction

mRNA molecules but the pool of available mRNA continues to be translated. In contrast, because of the direct regulation exerted by regulatory RNAs, sRNAs may destabilize the mRNA pool directly, achieving a faster regulation.

Second, sRNA regulation seems to achieve an inverse relation between the amount of free sRNA and free mRNA target (Figure 8B). The interaction of a transcription factor with the DNA usually involves one or a few molecules of transcription factors that regulate the synthesis of many molecules of mRNA. However, the interaction of a sRNA with its target mRNA may involve the interaction of one molecule of sRNA per one molecule of mRNA, achieving the inverse correlation between free sRNA and free mRNA target.

Third, this inverse correlation is used in some cases to achieve a threshold-linear response (Figure 8C). Some sRNA may be constitutively transcribed, leading to sequestration of every target mRNA molecule until the transcription rate of the target mRNA passes a certain threshold. However, when the transcription rate of the target mRNA overpasses the amount of sRNA, a linear response occurs that depends on the difference between the transcription rate of sRNA and target mRNA, a feature that could be used to filter weak or transient signals. All these features of sRNA regulation have been validated by several dynamic simulations (Shimoni et al., 2007; Mehta et al., 2008).

Finally, the combination of sRNA regulation and the regulation exerted by transcription factors may generate complex regulatory circuits. One of the simplest regulatory loops emerges when a transcription factors regulates the expression of a target mRNA and the expression of a sRNA that, in addition, regulates the same target mRNA (Mangan and Alon, 2003). These coherent or incoherent loops, depending on the logical relationships between their components, are thought to be essential, for example, in the filtering of spurious stimulus or the generation of pulses of expression (Alon, 2007). The overlay of several loops and layers of regulation generates an intricate network that fine-tunes the physiological responses. In fact, some analysis suggest that the post-transcriptional regulation exerted by sRNAs may play an important part (even more than classical transcriptional regulation) in the determination of the final concentration of gene products (Vogel and Marcotte, 2012).

22 Introduction

1.2.1 Types of RNA regulators

1.2.1.1 RNA regulators encoded in cis

RNA regulators that are transcribed from regions adjacent to the RNA molecule that they regulate are referred to as RNA regulators encoded in cis. The main group of cis encoded RNAs are asRNAs. These molecules are transcribed in antisense orientation to the mRNA target that they regulate, and, because of their genomic position, they show perfect, sometimes extensive, base complementarity to the coding region, 5’ UTR or 3’UTR of their mRNA target (Figure 7A). In some cases, the antisense regulation may be produced by the interaction of two mRNAs that regulate each other, a type of regulation referred to as “excludons” (Figure 7C-D) (Sesto et al., 2013; Georg and Hess, 2018). The mechanisms of asRNA regulation are based on the antisense orientation of the interacting molecules. asRNAs may interfere in the transcription of their mRNA targets by transcription interference or they may post-transcriptionally regulate the stability or translation efficiency of their mRNA targets (Figure 9) (Thomason and Storz, 2010; Georg and Hess, 2011; 2018).

Figure 9. Some asRNA regulatory mechanisms. The interaction by perfect base complementarity between an asRNA (red) and its target mRNA (blue) may provoke termination of transcription (A), mRNA degradation by RNases (orange) (B) or changes in the translation efficiency, generally by a block in the translation initiation mediated by interaction of the asRNA with the region containing the Shine-Dalgarno sequence (green) that prevents binding of the ribosome (grey) (C).

Antisense transcription was first observed in the context of mechanisms controlling plasmid copy number or associated to transposons, phages or toxin-antitoxin systems (Wagner and Simons, 1994). Nowadays, the use of transcriptomic data reveals that antisense

23 Introduction

transcription is prominent in all analysed bacteria and covers a high number of genes (Georg and Hess, 2018). In many cases, antisense transcription covers such a significant part of the genome that the term “pervasive transcription” has been adopted (Lybecker et al., 2014a). Some studies have shown that a basal antisense transcription provokes the degradation of the duplex of mRNA-asRNA mediated by RNase III (Lasa et al., 2011; Lybecker et al., 2014b). The function of this basal pervasive transcription is not completely understood yet. In contrast, the expression of some asRNAs is tightly regulated, suggesting a function related to the physiological signal that triggers their expression. For example, GadY is expressed in stationary phase (Opdyke et al., 2004) or SymR is expressed after DNA damage (Kawano et al., 2007) in E. coli.

Besides asRNAs, riboswitches are the other main group of cis encoded RNAs. They are usually part of the 5’ UTR of the bacterial mRNAs that they regulate. Riboswitches usually have two distinct functional domains, an “aptamer domain” that recognizes the effector molecules, and an “expression platform” that changes its structure in response to the binding of the effector to the aptamer domain and interacts with the transcriptional or translational machinery (Serganov and Nudler, 2013). Changes in the concentration of the ligands provoke conformational changes in the riboswitches that affect the transcription or translation of the mRNA target (Garst et al., 2011). Finally, in some cases, the stop of the transcription exerted by the riboswitch can generate a sRNA that may eventually act in trans (Loh et al., 2009).

1.2.1.2 RNA regulators encoded in trans

RNA regulators encoded in trans are transcribed from a different genomic region than their mRNA target(s). As it was mentioned before, some sRNAs are transcribed from their own genes located in intergenic regions (Figure 7D) but others are transcribed from promoters located in the 5’ UTR, coding region or 3’UTR of another gene (Figure 7F). In addition, some sRNAs may be generated from the processing of a longer mRNA molecule (Figure 7E) (Miyakoshi et al., 2015).

In contrast to asRNAs, that interact by perfect base complementarity with their targets, the interaction between sRNAs and their targets takes place by short and imperfect base pairing (Bobrovskyy and Vanderpool, 2013). The interaction region usually comprises 10 to 25 nucleotides (Peer and Margalit, 2011). Some of these nucleotides are highly conserved for each given sRNA and form what is called the “seed” region (Storz et al., 2011), a region involved in the initial recognition of the target mRNA. This seed region usually consists of 6 or 7 nucleotides, is usually located near the 5’ end of the sRNA (Guillier and Gottesman, 2008; Papenfort et al.,

24 Introduction

2010) and seems to be essential for the specificity of the regulation because one single mutation in this region can abolish the specificity of the regulation exerted by some sRNAs (Papenfort et al., 2012). In addition, most sRNAs have a Rho-independent terminator, a strong secondary structure followed by multiple uracils (polyU) that favours the termination of transcription and protects the sRNA from 3’-exonucleases.

Small RNAs may positively or negatively regulate the expression of their target mRNAs by diverse mechanisms (Figure 10).

Figure 10. Some sRNA regulatory mechanisms. sRNAs encoded in trans (red), here transcribed from an intergenic region, interact with their target mRNAs (blue) by imperfect base pairing. (A) The binding of the sRNA at the translation start region of the target mRNA, including the ribosome binding site (green), prevents the binding of the ribosome (grey) and, consequently, translation initiation is blocked. (B) The duplex sRNA-mRNA is recognized by a RNase (orange) and mRNA may be degraded. (C) sRNA may also exert a positive regulation in the translation efficiency of a mRNA. The binding of the sRNA may provoke a conformational change that may release the sequestered Shine- Dalgarno sequence, improving the translation initiation.

A very common mechanism involves the binding of the sRNA to the 5’UTR of its target mRNA in a region that overlaps or is located near to the ribosome binding site (RBS). The binding of the sRNA usually blocks the translation (Figure 10A) and, in many cases, promotes the degradation of the mRNA by the recruitment of different RNases, such as RNase E or RNase III (Figure 10B) (Wagner and Romby, 2015). However, sRNAs may also exert a positive regulation. The RBS of an mRNA may be sequestered by the strong secondary structure of its own 5’UTR. In these cases, the binding of an sRNA to the 5’UTR may provoke a conformational change that

25 Introduction

releases the RBS and favours the translation (Figure 10C) (Fröhlich and Vogel, 2009). In addition, some sRNA regulate the expression of a specific gene that is transcribed in a polycistronic transcript. In these cases, the regulation exerted by the sRNA leads to uncoupling of the expression of the genes encoded in the operon, as in the regulation of the galactose operon galETKM by the sRNA Spot42 in E. coli (Møller et al., 2002).

Although the binding of the sRNAs to the 5’UTR of their mRNA targets is the most common mechanism, sRNAs can exert their function through a variety of alternative mechanisms. They may bind inside the coding region or in the 3’ UTR (Jagodnik et al., 2017). In addition, they may also interact with RNA-binding proteins. For example, some sRNAs have sequences that mimic the binding sites of protein regulators and can transiently sequester the protein regulator. A well described example is the regulation of carbon usage in E. coli exerted by the sRNAs CsrB and CsrC and the protein regulator CsrA (Liu et al., 1997). In this conserved regulatory circuit, when CsrB and CsrC are expressed under low-nutrient conditions CsrA is sequestered by the multiple binding sites present in these two sRNAs and, consequently, is unable to regulate its targets (Babitzke and Romeo, 2007).

The function of many sRNAs depends on the RNA chaperone Hfq. The role of this protein is not very relevant in the mechanisms operated by cis encoded RNAs (Brantl, 2007) but in many bacteria it is necessary for the stabilization of many sRNAs and is required for the short and imperfect interaction of many sRNAs with their targets (Updegrove et al., 2016). Hfq is not conserved in all bacterial genomes (Sun et al., 2002) but recent reports have identified new proteins with a similar role (Attaiech et al., 2017) such as YbeY, a protein with a MID domain of the eukaryotic Argonaute proteins (Pandey et al., 2011), and ProQ, a RNA chaperone with a FinO domain (Smirnov et al., 2016), a conserved domain that recognizes structural motifs of the sRNAs (Holmqvist and Vogel, 2018).

A homolog of Hfq is encoded in many cyanobacterial genomes. However, although the crystal structure of Hfq from Nostoc sp. PCC 7120 and Synechocystis sp. PCC 6803 showed a structure similar to that of other bacterial Hfq proteins, cyanobacterial Hfq has low affinity for well-known sRNAs of E. coli in vitro, is unable to mediate the regulation exerted by sRNAs in vivo in E. coli and shows some differences in the binding sites of RNAs (Bøggild et al., 2009). It has been described that the lack of Hfq affects nitrate assimilation in Nostoc sp. PCC 7120 (Puerta- Fernández and Vioque, 2011) or motility in Synechocystis sp. PCC 6803 (Dienst et al., 2008), in this last case through the involvement of Hfq in protein-protein interactions rather than protein- RNA interactions (Schuergers et al., 2014).

26 Introduction

1.2.2 Identification of RNA regulators and their targets

Non-coding RNAs have been largely overlooked because they are difficult to detect and characterise. Some conserved non-coding RNAs (as tmRNA, the RNA component of RNase P, 4.5S and 6S RNAs) were first discovered by chance because of its abundance or its role in housekeeping functions. Only ten non-coding RNAs were known in E. coli in 2001 (Wassarman et al., 1999). However, the availability of genomic information allowed the design of bioinformatic approaches for the identification of new non-coding RNAs (Altuvia, 2007). Because sRNAs share some features that allow the design of global approaches for their discovery, some predictive algorithms were developed before transcriptomic data were available. First global approaches were based on the prediction of sequences in intergenic regions flanked by transcription signals and putative Rho-independent terminators (Argaman et al., 2001; Rivas et al., 2001; Wassarman et al., 2001; Chen et al., 2002). The conservation of the primary sequence (Argaman et al., 2001; Wassarman et al., 2001), putative RNA structural domains (Rivas et al., 2001) or both (Washietl et al., 2005) (Livny et al., 2006) were taken as indications pointing to the existence of a gene transcribed as an sRNA. In addition, the use of microarrays with probes for intergenic regions allowed the identification of sRNAs that were not widely conserved (Tjaden et al., 2002; Zhang et al., 2003). Later, the availability of transcriptomic data based on RNAseq allowed the identification of not only sRNAs but also asRNAs (Sorek and Cossart, 2010). Among the variants of RNAseq, differential RNAseq (dRNAseq) has been a successful technique for the discovery of new RNA regulators (Sharma and Vogel, 2014). This methodology is based on the pre-treatment of the RNA samples with TEX exonuclease (Terminator 5’-Phosphate-dependent Exonuclease), an RNase that degrades transcripts with 5’-monophosphate, so that the resulting RNA samples are enriched in primary transcripts with 5’-triphosphate, allowing the precise identification of the TSS. This methodology had been applied in Nostoc sp. PCC 7120 (Mitschke et al., 2011b) and the resulting dataset has been fundamental for the computational approach that is presented in this Thesis.

cis encoded RNAs, such as asRNAs and riboswitches, are transcribed in the same locus than the mRNA they regulate, although some asRNAs or riboswitches may additionally act as sRNAs (Loh et al., 2009; Georg and Hess, 2018). However, the identification of target mRNAs regulated by sRNAs is not a trivial task. The phenotypes associated to lack of sRNAs are usually subtle. In addition, because the interactions of sRNAs with their target mRNAs are short and imperfect, the computational prediction of targets is difficult. CopraRNA is one of the most advanced algorithms used for the prediction of mRNA targets (Wright et al., 2013; Wright et al.,

27 Introduction

2014). This algorithm searches through the predicted transcriptome of one organism to identify putative targets of a sRNA based on sequence complementarity, folding energy and hybridization energy. However, it also implements a phylogenetic criterion that considers the conservation of any predicted interaction across several species and the emergence of compensatory mutations that support the occurrence of the putative interaction in vivo.

Recent high throughput methodologies allow the direct identification of targets in vivo. MAPS (MS2-affinity purification coupled with RNA sequencing) is a methodology based on the co-immunopurification of an sRNA tagged with the aptamer for the protein of MS2 phage capsid and its target mRNAs, followed by an RNAseq sequencing protocol (Lalaouna et al., 2017). GRIL- Seq (Global sRNA Target Identification by Ligation and Sequencing) is based on the preferential ligation by T4 RNA ligase of sRNAs with the ends of their interacting target mRNAs, followed by RNAseq sequencing and the identification of the chimaeras (Han et al., 2016).The use of these two methodologies has allowed the definition of the “targetome” of some sRNAs (Lalaouna et al., 2015a; Lalaouna et al., 2015b; Zhang et al., 2017; Lalaouna et al., 2018).

1.2.3 RNA regulators in cyanobacteria

1.2.3.1 Identification of RNA regulators in cyanobacteria

Global approaches for the identification of cyanobacterial RNA regulators have been based on the use of computational tools followed by experimental validation (Axmann et al., 2005; Voss et al., 2009; Ionescu et al., 2010) and microarray analysis (Steglich et al., 2008; Georg et al., 2009; Gierga et al., 2012). In addition, transcriptomic data based on RNAseq in model unicellular cyanobacteria such as Synechocystis sp. PCC 6714 (Kopf et al., 2014a; Kopf et al., 2014c), Synechocystis sp. PCC 6803 (Mitschke et al., 2011a; Billis et al., 2014; Kopf et al., 2014b) or Synechococcus elongatus sp. PCC 7942 (Vijayan et al., 2011; Billis et al., 2014), relevant species in natural environments such as Trichodesmium erythraeum (Pfreundt et al., 2014), Nodularia spumigena CCY9414 (Voss et al., 2013; Kopf et al., 2015b), marine Synechococcus (Gierga et al., 2012), or Prochlorococcus (Steglich et al., 2008; Thompson et al., 2011; Waldbauer et al., 2012; Voigt et al., 2014), and heterocyst-forming species such as Nostoc sp. PCC 7120 (Flaherty et al., 2011; Mitschke et al., 2011b), have revealed that non-coding transcripts are associated to more than 10 % of active promoters.

A transcriptomic analysis of Nostoc sp. PCC 7120 was carried by our group using a dRNAseq methodology (Mitschke et al., 2011b). In this approach, the transcriptional responses

28 Introduction to nitrogen deficiency of a wild-type (WT) and a hetR mutant strains were studied. The comparison between transcriptional changes that took place in both strains (general response to nitrogen deficiency, DEF) versus only in the WT strain (HetR-dependent responses specifically involved in heterocyst differentiation, DIF) allowed the dissection of two categories of genes, DEF and DIF groups respectively (Mitschke et al., 2011b). Both categories contained many TSS that could be associated to asRNAs or sRNAs. This dataset has been extensively used in the approaches included in this Thesis. The precise determination of TSS in intergenic regions has allowed the design of microarrays with probes associated to sRNAs and asRNAs and the implementation of a bioinformatic pipeline for the identification of new sRNAs.

1.2.3.2 Physiological processes regulated by RNA regulators in cyanobacteria

Although our knowledge of the physiological roles of asRNAs and sRNAs in some bacteria, in particular enterobacteria, has increased exponentially in recent years, the function of most cyanobacterial asRNAs and sRNAs remains unknown (Kopf and Hess, 2015). One of the first identified sRNAs in cyanobacteria was Yfr1, a sRNA discovered in a global approach in Phrochlorococcus (Axmann et al., 2005). Yfr1 is conserved in all cyanobacterial genomes analyzed (Voss et al., 2007), is required for proper growth under multiple stresses in Synechococcus elongatus (Nakamura et al., 2007) and may interact with some mRNAs encoding porins (Richter et al., 2010). Its mechanism of regulation or the physiological process in which it may be involved remain unclear, although recently it has been shown that it might be regulated by interaction with another conserved sRNA, Yfr2 (Lambrecht et al., 2019).

It is not surprising that some of the asRNAs and sRNAs already described in cyanobacteria play a role in the post-transcriptional regulation of the photosynthetic process (Wilde and Hihara, 2016). PsrR1 (photosynthesis regulatory RNA 1) is a well conserved RNA first discovered by a computational approach carried out in Synechocystis sp. PCC 6803 (Voss et al., 2009). Its expression increases under high light conditions (Georg et al., 2009; Mitschke et al., 2011a) and is subjected to daily cycles with a peak early in the morning (Beck et al., 2014). Computational analysis predicts that PsrR1 may interact with the region near the RBS of many mRNAs that encode structural proteins of the photosynthetic apparatus. As an example, PsrR1 interacts with psaL mRNA, that encodes subunit IX of the reaction center of PSI. PsrR1 blocks its translation and favours its degradation by RNase E (Georg et al., 2014). In addition, the expression of PsrR1 is regulated by RpaB (Kadowaki et al., 2016), a transcriptional regulator that regulates acclimation to high light. So, because RpaB regulates the expression of PsrR1 and psaL,

29 Introduction

RpaB and PsrR1 are part of a regulatory coherent loop that regulates acclimation to changing light conditions. Ncr0700/PmgR1 (photomixotrophic growth RNA 1) is another sRNA with an expression profile opposite to PsrR1. PmgR1 accumulates in the dark (Beck et al., 2014), regulates the accumulation of glycogen and is essential for mixotrophic growth (de Porcellinis et al., 2016).

Photosynthesis is also regulated by asRNAs in Synechocystis sp. PCC 6803. The psbA2 and psbA3 mRNAs, encoding the D1 protein of the reaction center of PSII, are degraded by RNase E in the dark at the AU-rich motifs located in the 5’ UTRs of these mRNAs (Sakurai et al., 2012). The maintenance of the levels of psbA mRNAs in the light is achieved through a positive regulation exerted by two asRNAs complementary to the 5’UTR of these mRNAs. In the light, the asRNAs are upregulated, interact with the AU-rich motifs and prevent the cleavage of the mRNAs by RNase E (Sakurai et al., 2012). Flv4 and Flv2 are flavodiiron proteins that protect PSII against oxidative stress (Zhang et al., 2012) and IsiA is a chlorophyll binding protein that, under iron deficiency, forms an additional antenna around PSI (Chauhan et al., 2011). The expression of the operon flv4-2 and the isiA mRNA are negatively regulated by the asRNAs As1_flv4 (Eisenhut et al., 2012) and IsrR (Dühring et al., 2006), respectively. In both cases, the constitutive expression of the asRNAs and the co-degradation with their target mRNAs establish a certain threshold for the protein synthesis (Legewie et al., 2008), ensuring that Flv4 and Flv2 or IsiA are

only translated under low CO2 conditions or iron deficiency, respectively.

Some asRNAs and sRNAs have also been described in the heterocyst-forming strain Nostoc sp. PCC 7120. One of the first asRNA described in cyanobacteria was an antisense complementary to furA (all1691). This antisense is produced from the transcription of alr1690, an adjacent gene in antisense orientation, it partially covers the sequences encoding FurA and post-transcriptionally regulates the expression of this transcriptional regulator involved in iron metabolism (Hernández et al., 2006).

Because the adaptation to nitrogen deprivation in Nostoc sp. PCC 7120 is a well- studied process, most sRNAs described in this organism so far are regulated by nitrogen availability. Some sRNAs, as NsiR3 (Mitschke et al., 2011b), NsiR4 (Klähn et al., 2015) or NsrR1 (Álvarez-Escribano et al., 2018), may be involved in the general response to nitrogen deprivation whereas transcription of other sRNAs, such as NsiR1 (Ionescu et al., 2010) is specifically induced in heterocyst, suggesting it might participate in the regulation of heterocyst differentiation. NsiR1 is transcribed from multiple copies in the genome of heterocyst-forming cyanobacteria

30 Introduction

(Ionescu et al., 2010) and is expressed at a very early stage in the differentiation process (Muro- Pastor, 2014) but its function has remained elusive.

NsiR4 (nitrogen stress-induced RNA 4) is a sRNA that participates in the regulation of the assimilation of nitrogen through the post-transcriptional regulation of IF7, one of the inactivating factors of glutamine synthetase in Synechocystis sp. PCC 6803 (Klähn et al., 2015). Because the expression of NsiR4 and IF7 are regulated by NtcA, NsiR4 is involved in a coherent regulatory loop that regulates one of the key enzymes in the assimilation of nitrogen (Klähn et al., 2015).

NsrR1 (nitrogen stress-repressed RNA 1) is a sRNA directly repressed by NtcA under nitrogen deprivation and interacts with the 5’UTR of the nblA mRNA promoting its degradation (Álvarez-Escribano et al., 2018). As it is the case of NsiR4, because NtcA also regulates the expression of nblA, NsrR1, NtcA and NblA also form a coherent regulatory loop that regulates the recycling of phycobilisomes under changes in nitrogen availability (Álvarez-Escribano et al., 2018).

Some sRNAs and asRNAs are specifically induced in heterocyst and could be involved in the regulation of heterocyst differentiation. A recent report identified an asRNA (As_glpX) specifically induced in heterocysts that regulates the degradation of glpX mRNA (Olmedo-Verd et al., 2019). glpX encodes sedoheptulose-1,7-bisphosphatase/fructose-1,6-bisphosphatase (SBPase), a key enzyme in the Calvin cycle. Thus, this asRNA contributes to the metabolic reprogramming of heterocyst by locally diminishing the levels of one of the enzymes necessary for CO2 fixation.

The main goal of this Thesis has been the identification and characterization of asRNAs and sRNAs in Nostoc sp. PCC 7120 that may be involved in the regulation of heterocyst differentiation or in general physiological process that may affect this differentiation process.

31

2. OBJECTIVES

Objectives

2. OBJECTIVES

The aim of this Thesis is the identification and characterization of sRNAs and asRNAs that may be involved in the regulation of the responses to nitrogen deprivation in Nostoc sp. PCC 7120. The following specific objectives are addressed:

1. Identification of new sRNAs by a bioinformatic approach. 2. Functional characterization of Yfr1, a phylogenetically conserved sRNA. 3. Generation of a co-expression network, which includes non-coding RNAs, to dissect the complex changes in the transcriptome taking place during heterocyst differentiation. 4. Functional characterization of NsiR1, a sRNA specifically induced in heterocysts.

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3. SUMMARY OF RESULTS

Summary of results

3. SUMMARY OF RESULTS

Regulation of gene expression cannot be understood without the participation of RNA regulators. Because these molecules are difficult to detect and usually exert a subtle post- transcriptional regulation, their functions have remained elusive for many years. The emergence of “omic” technologies and the use of global approaches have increased the number of new RNA regulators identified. The global objective of this Thesis is the use of computational approaches for the identification of new sRNAs and asRNAs in the heterocyst-forming cyanobacterium Nostoc sp. PCC 7120 and the functional characterization of some of those RNAs. The content of this dissertation is structured in four chapters, two of them already published as research articles.

3.1 Chapter I: A computational approach for the identification of conserved sRNAs in heterocyst-forming cyanobacteria.

Brenes-Álvarez, M., Olmedo-Verd, E., Vioque, A., & Muro-Pastor, A. M. (2016). Identification of conserved and potentially regulatory small RNAs in heterocystous cyanobacteria. Front Microbiol, 7, 48. doi:10.3389/fmicb.2016.00048

Previous computational approaches to the identification of sRNAs had been focused in the conservation of sequences located in intergenic regions flanked by predicted transcriptional signals (-10, -35 boxes) and putative Rho-independent terminators. Whereas the software for the prediction of Rho-independent terminators (a stem loop followed by a poly uracil stretch) is quite accurate, the prediction of promoter motifs and other transcriptional signals is not very accurate. In our work, we overcome this drawback by using an experimental dataset from a previous dRNAseq study that determined most of the TSS in Nostoc sp. PCC 7120 (Mitschke et al., 2011b). The use of this dataset together with sequence information from 89 cyanobacterial genomes allowed us to predict 327 putative sRNAs.

Basically, we searched in the genome of Nostoc sp. PCC 7120 for sequences located in intergenic regions that were flanked by a TSS and a Rho-independent terminator and with a size range of 45 - 350 nucleotides. Sequences that fulfilled those criteria were extracted and

39 Summary of results

compared to a database of 89 complete cyanobacterial genomes. The output was a dataset of potential sRNAs with different degrees of phylogenetic conservation.

Validation by Northern blot of transcription of new predicted sRNAs and the presence of previously validated sRNAs in our dataset demonstrated the performance of our approach. Finally, our phylogenetic analysis showed widely conserved sRNAs, such as Yfr1 (see Chapter II), but also some sRNAs only conserved in heterocystous cyanobacteria, which may suggest a putative function of these sRNAs in one of the processes that are exclusive of these cyanobacteria, the differentiation of heterocysts. In fact, the heterocyst-specific transcription of one of these sRNAs (NsiR8) was also validated.

3.2 Chapter II: Yfr1, a widely conserved sRNA, regulates the integrity of the cell wall and its remodeling during heterocyst differentiation.

Brenes-Álvarez, M., Vioque, A., & Muro-Pastor, A. M. (2019). Integrity of the cell wall and its remodeling during heterocyst differentiation is regulated by phylogenetically conserved sRNA Yfr1 in Nostoc sp. PCC 7120. (Submitted).

Yfr1 was one of the first sRNAs described in cyanobacteria (Axmann et al., 2005). A previous study (Voss et al., 2007) and our computational approach (see Chapter I) had shown that Yfr1 is conserved in all available cyanobacterial genomes, pointing to a role in the regulation of an essential physiological process. So, we used some of the homologs detected in our computational approach to predict putative mRNA targets of Yfr1.

The prediction of targets showed an enrichment in genes encoding outer membrane proteins or enzymes related to the biosynthesis and remodeling of peptidoglycan. Some of the predicted targets were validated using a heterologous reporter system in E. coli. Nostoc sp. PCC 7120 strains with altered levels of Yfr1 were generated and the expression of murB and mraY, two genes that encode enzymes that participate in the synthesis of peptidoglycan, was tested in those strains, confirming the previously validated interactions.

The phenotypic characterization of the strains with altered levels of Yfr1 showed that they are affected in the integrity of cell envelopes. They had a higher sensitivity to harmful substances that should not pass the normal cyanobacterial envelopes. They were also sensitive

40 Summary of results to vancomycin, an antibiotic that specifically binds to the nascent chains of peptidoglycan, and they showed an abnormal distribution of nascent septa.

In addition, the strain that overexpressed Yfr1 showed a poor diazotrophic growth. A close inspection showed that filaments of this strain growing in media without combined nitrogen were usually broken at the connections between vegetative cells and heterocysts. Thus, these results additionally confirmed the importance of peptidoglycan synthesis and remodeling during the differentiation of heterocyst.

3.3 Chapter III: A co-expression network to dissect the complex transcriptome of Nostoc sp. PCC 7120 during heterocyst differentiation.

Brenes-Álvarez, M., Mitschke, J., Olmedo-Verd, E., Georg, J., Hess, W.R., Vioque, A., and Muro-Pastor, A.M. (2019). Elements of the heterocyst-specific transcriptome unravelled by co-expression analysis in Nostoc sp. PCC 7120. Environmental Microbiology 21, 2544-2558. doi: 10.1111/1462-2920.14647.

When filaments of Nostoc sp. PCC 7120 are growing in media without combined nitrogen, the filaments are formed by a combination of vegetative cells, mature N2-fixing heterocysts and immature heterocysts. The analysis of transcriptomes in these different cell types is not possible because they can´t be physically isolated with their RNA intact. Thus, we have followed a genetic strategy and the generation of a co-expression network in order to dissect the complex transcriptome of our model organism.

A co-expression network is a graphical representation in which elements with a similar expression profile are grouped together. Because the aim of this Thesis was the study of the regulation exerted by asRNAs and sRNAs, one of the features of our co-expression network was the inclusion of not only protein-coding genes but also non-coding RNAs, asRNAs and sRNAs. Customized microarrays were designed based on the previous RNAseq study that had determined TSS associated to genes, asRNAs and sRNAs (Mitschke et al., 2011b). These microarrays were hybridized with RNA samples extracted from the wild-type and the hetR mutant strain at different times after removal of combined nitrogen.

The use of the co-expression network and a clustering analysis allowed the identification of groups of elements (genes, asRNAs and sRNAs) with similar expression profiles.

41 Summary of results

Some of these groups were related to the general response to nitrogen deprivation that occurs in all cells of the filament, commented in section 1.1.4.1. In contrast, other groups could be specifically associated to heterocyst differentiation. So, the combination of the co-expression network and the clustering analysis allowed us to classify groups of elements based on their spatial (all cells or only heterocysts) or temporal (early or late in the differentiation process) expression. In addition, the comparison of the promoter sequences of genes (and its homologs) that have been found in the same group showed an enrichment in two motifs (DIF1 and DIF2) that were linked to cell-specific transcription in heterocysts.

Many previously described genes involved in heterocyst differentiation were found in the groups specifically linked to differentiation, confirming the validity of our approach. In addition, we also found new sRNAs and asRNAs among the elements included in these groups. The cell-specific expression of one asRNA (as_alr5059) and one sRNA (NsiR9) was validated. So, these results highlight the putative regulation exerted by non-coding RNAs during the process of heterocyst differentiation.

3.4 Chapter IV: NsiR1, a sRNA with multiple copies, regulates heterocyst differentiation.

Brenes-Álvarez, M., Minguet, M., Vioque, A., & Muro-Pastor, A. M. (2019). NsiR1, a small RNA with multiple copies, regulates heterocyst differentiation and commitment in Nostoc sp. PCC 7120. (In preparation).

NsiR1 is a sRNA encoded in 12 tandem copies in the region upstream to hetF in the genome of Nostoc sp. PCC 7120 and other heterocyst-forming cyanobacteria (Ionescu et al., 2010). The seven central copies (#3-#9) are identical whereas the rest of the copies show some sequence divergence. The different copies are transcribed from individual promoters that contain a DIF1 motif (Ionescu et al., 2010), one of the DNA motifs that was linked to an early specific transcription in cells becoming heterocyst according to our co-expression network and clustering analysis (Brenes-Álvarez et al., 2019). NsiR1 is expressed in cells becoming heterocyst before any sign of morphological differentiation is observed (Muro-Pastor, 2014). In addition, based on transcriptomic information (Mitschke et al., 2011b) and the genomic context, the copy

42 Summary of results that is located closest to hetF (NsiR1.1) may act as an antisense complementary to the 5’UTR of hetF mRNA.

The interaction between NsiR1.1 and hetF was validated using an in vivo reporter system in E. coli. NsiR1.1 seems to negatively regulate the expression of hetF and in vitro analysis showed that this negative regulation may be achieved through a conformational change that reduces the accessibility of the translation machinery to the region next to the start codon.

Our phylogeny analysis showed that the high number of tandem copies is conserved in genomes of heterocystous cyanobacteria. The putative functional divergence between NsiR1.1 and NsiR1.4 was tested through the generation of strains that overexpressed NsiR1.1 (OE_NsiR1.1), NsiR1.4 (OE_NsiR1.4) or a sponge that depleted the pool of NsiR1 RNA (OE_as_NsiR1). When we analyzed the phenotype of these strains, the overexpression of NsiR1.1 resulted in higher levels of HetR. This is the expected result if NsiR1 reduces the expression of HetF, as HetF is a protease that degrades HetR (Risser and Callahan, 2008).

The phenotypic characterization of the Nostoc strains showed that filaments of OE_NsiR1.4 strain contained heterocysts when they were grown in media with nitrate, a condition in which heterocyst differentiation should not take place. This phenotype was not observed in the OE_NsiR1.1 strain. As a consequence, a functional divergence between different copies of NsiR1 was demonstrated. A prediction of targets for NsiR1.4 had detected a possible interaction between NsiR1.4 and alr3234 5’UTR (Ionescu et al., 2010), a gene related to heterocyst-commitment to differentiation. We have confirmed such interaction. Thus, the diversification of the copies of NsiR1 allows the cell-specific post-transcriptional regulation of two processes involved in the differentiation of heterocysts.

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3.1 CHAPTER I A computational approach for the identification of conserved sRNAs in heterocyst-forming cyanobacteria

CHAPTER I. A computational approach for the identification of conserved sRNAs in heterocyst-forming cyanobacteria

Brenes-Álvarez, M., Olmedo-Verd, E., Vioque, A., and Muro-Pastor, A.M. (2016). Identification of conserved and potentially regulatory small RNAs in heterocystous cyanobacteria. Frontiers in Microbiology 7, 48. doi: 10.3389/fmicb.2016.00048.

3.2 CHAPTER II Yfr1, a widely conserved sRNA, regulates the integrity of the cell wall and its remodeling during heterocyst differentiation

Integrity of the cell wall and its remodeling during heterocyst differentiation is regulated by phylogenetically conserved sRNA Yfr1 in Nostoc sp. PCC 7120

Manuel Brenes-Álvarez, Agustín Vioque and Alicia M. Muro-Pastor#

Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and Universidad de Sevilla, E-41092 Sevilla, Spain

#Corresponding author: Alicia M. Muro-Pastor, Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and Universidad de Sevilla, E-41092 Sevilla, Spain

Tel. +34-954489521

Email: [email protected] ORCID iD: 0000-0003-2503-6336

Running Title: Yfr1 sRNA regulation in Nostoc 7120

Word count (abstract): 233 Word count (text): 4892

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ABSTRACT

Yfr1 is a strictly conserved small RNA in Cyanobacteria. A bioinformatic prediction to identify possible interactions of Yfr1 with mRNAs was carried out using the sequences of Yfr1 from several heterocyst-forming strains, including Nostoc sp. PCC 7120. The results of the prediction were enriched in genes encoding outer membrane proteins and enzymes related to peptidoglycan biosynthesis and turnover. Heterologous expression assays in Escherichia coli demonstrated direct interactions of Yfr1 with mRNAs of eleven of the candidate genes. The expression of ten of them (alr2458, alr4550, murC, all4829, all2158, mraY, alr2269, alr0834, conR, patN) was repressed by interaction with Yfr1, whereas the expression of amiC2, encoding an amidase, was increased. The interactions between Yfr1 and the eleven mRNAs were confirmed by site-directed mutagenesis of Yfr1. Furthermore, a Nostoc strain with reduced levels of Yfr1 had higher amounts of mraY and murC mRNAs, supporting a role for Yfr1 in the regulation of those genes. Nostoc strains with either reduced or increased expression of Yfr1 showed anomalies in cell wall completion and were more sensitive to vancomycin than the wild type strain. Furthermore, growth in the absence of combined nitrogen, that involves differentiation of heterocysts, was compromised in the strain overexpressing Yfr1, and filaments were broken at the connections between vegetative cells and heterocysts. These results indicate that Yfr1 is an important regulator of cell wall homeostasis and correct cell wall remodeling during heterocyst differentiation.

IMPORTANCE

Bacterial small RNAs (sRNAs) are important players affecting the regulation of essentially every aspect of bacterial physiology. The cell wall is a highly dynamic structure that protects bacteria from their fluctuating environment. Cell envelope remodeling is particularly critical for bacteria that undergo differentiation processes such as spore formation or differentiation of heterocysts. Heterocyst development involves the deposition of additional layers of glycolipids and polysaccharides outside the outer membrane. Here we show that a cyanobacterial phylogenetically conserved small regulatory RNA, Yfr1, coordinates expression of proteins involved in cell wall-related processes, including peptidoglycan metabolism and transport of different molecules, as well as expression of several proteins involved in heterocyst differentiation.

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INTRODUCTION

Bacterial cell envelopes are multi-layered structures that delimit the interior of the cell from its environment. The essential component of bacterial cell walls is peptidoglycan, a strong and flexible mesh that protects the cell against osmotic pressure and contributes to the shape of the cell. According to the architecture of the envelopes, bacteria are classified in two groups. Gram-positive strains have a thick peptidoglycan layer (30 to 100 nm) surrounding the cytoplasmic membrane, while Gram-negative strains have a thin peptidoglycan layer (only a few nm) between the inner, cytoplasmic membrane, and a second, outer membrane (1). Cyanobacteria are Gram-negative. However, the peptidoglycan layer between the inner and outer membranes is relatively thick (15-30 nm in filamentous strains) and characterized by extensive crosslinking, rather resembling the architecture of Gram-positive bacteria (2).

The sacculus of peptidoglycan is a dynamic structure that must adapt to the growth of the cells, the separation of daughter cells during cell division, the turnover of peptidoglycan or the assembly of large trans-envelope complexes (e. g. secretion systems). In addition, cellular differentiation processes that affect cell envelopes involve peptidoglycan remodeling, performed by different murein hydrolases, such as N-Acetylmuramoyl-L-Ala amidases (Ami enzymes), reviewed in (3).

Nostoc sp. PCC 7120 is a filamentous cyanobacterium that under nitrogen deprivation differentiates heterocysts (specialized cells devoted to N2 fixation) in a semiregular pattern (4). Heterocyst differentiation involves biochemical and morphological changes that provide a micro-oxic environment for nitrogenase, an O2-labile enzyme. In addition to the inactivation of the O2-producing photosystem II and the increase in the O2 consumption rate, heterocysts have a special cellular envelope with two extra layers, an external one composed of polysaccharides (HEPs) and a laminated, internal layer composed of glycolipids (HGLs) that acts as a barrier to gas diffusion (5). Both heterocyst-specific layers are deposited outside the outer membrane of these cells.

Under nitrogen-fixing conditions there is a metabolic division of labor between vegetative cells and heterocysts in cyanobacterial filaments. Heterocysts feed vegetative cells with fixed nitrogen and obtain fixed carbon in return. Growth of nitrogen-fixing filaments depends on the transport and exchange of metabolites between vegetative cells and heterocysts (4). One route for this exchange of metabolites could consist of diffusion from cytoplasm to cytoplasm through septal junctions, protein structures that allow the

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intercommunication between the cytoplasms of adjacent cells (6-9). These structures traverse the septal peptidoglycan disks through perforations called “nanopores” (7) that may be made by murein hydrolases.

Mutants in murein amidases AmiC1 (alr0092) (10) and AmiC2 (alr0093, hcwA) (11, 12) show a significant reduction in the number of nanopores and defects in diazotrophic growth. In addition, mutants in peptidoglycan synthesis enzymes MurC (alr5065) and MurB (alr5066) (13) and in some penicillin binding proteins (all2981, alr4579 and alr5051) (14, 15) show alterations in heterocyst differentiation and growth defects in media without combined nitrogen. All this evidence points to synthesis and remodeling of peptidoglycan being essential for proper heterocyst differentiation and diazotrophic growth. In fact, several genes related to peptidoglycan metabolism are transcribed from HetR-dependent, heterocyst-specific promoters (16, 17).

The maintenance and assembly of the outer membrane relies on proteins of the Omp85 family. Omp85 is essential for outer membrane maturation in E.coli (18) and a mutant of alr2269 (encoding a Omp85 homolog in Nostoc sp. PCC 7120) showed a disturbed outer membrane and, as a consequence, higher sensitivity to harmful substances, such as erythromycin (19). Molecules enter the periplasm through different transporters, such as porin- like proteins and TonB-dependent transporters. Porins usually allow the diffusion of hydrophilic molecules of a size up to 600 Da with low selectivity (20). The genome of Nostoc sp. PCC 7120 encodes a general porin (Alr0834), OprB-like porins (Alr4550 and All4499) (21), and several TonB-dependent transporters (All2158, All3310, All4026) (22).

Yfr1 is one of the first small RNAs (sRNA) identified in Cyanobacteria and was initially described in unicellular picocyanobacteria (23). Bioinformatic prediction and the use of a heterologous reporter system in E.coli demonstrated that Yfr1 can interact with the mRNAs encoding two porins in Prochlorococcus MED4 (24). Recently, a global approach allowed the identification of the targetome of Yfr1 from Prochlorococcus MED4 (25). In Synechococcus elongatus PCC 6301, Yfr1 is highly expressed, with slight abundance changes in cells exposed to high salt stress or oxidative stress. A Yfr1 mutant showed reduced growth under iron limitation, high salt stress or oxidative stress (26).

Yfr1 has been identified in all cyanobacterial genomes analyzed, from the minimal genomes of unicellular strains of the Prochlorococcus-Synechococcus lineages to much larger genomes of complex filamentous strains, such as Nostoc, able to undergo cellular differentiation processes (27). This broad occurrence suggests a widely conserved function for Yfr1.

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Interestingly, a fully conserved sequence motif is present in all Yfr1 homologs across the different cyanobacterial clades (27, 28). We have identified and verified the interaction of Yfr1 with several mRNAs whose products are involved in peptidoglycan metabolism and envelope biogenesis and maintenance in Nostoc sp. PCC 7120, including several proteins required for proper differentiation of functional heterocysts and therefore N2 fixation. Our results suggest that Yfr1 may regulate the composition and remodeling of envelopes of heterocyst-forming cyanobacteria.

RESULTS

Validation of putative Yfr1 targets in E. coli.

According to previous studies, Yfr1 is expressed constitutively, with only slight expression changes under different growth conditions in the unicellular cyanobacteria Synechococcus elongatus PCC 6301 (26) or Prochlorococcus MED4 (25). We tested expression of Yfr1 in Nostoc sp. PCC 7120 (Fig. S1). Similar to the observations made in the case of unicellular strains, Yfr1 had a relatively strong expression and its levels changed only slightly due to nitrogen deficiency or high light stress, two conditions that lead to pronounced changes in cyanobacterial gene expression. As previously shown, Yfr1 accumulates in the form of two transcripts with slightly different size (28).

The sequence of Yfr1 from Nostoc sp. PCC 7120 is shown in Fig. 1A. Because prediction of sRNA targets can be improved when comparative phylogenetic information is taken into account (29), we used CopraRNA, a software to predict potential interactions between sRNAs and mRNA targets that are conserved among a set of organisms (29, 30). Using the sequences of Yfr1 homologs from ten different heterocystous strains (see Materials and Methods for details), the resulting list of predicted targets (Table S1) showed a significant enrichment in transporters, enzymes related to cell wall synthesis or remodeling and proteins located in the outer membrane. Amongst the predicted targets were several mRNAs encoding enzymes involved in peptidoglycan metabolism such as alr2458 (alanine racemase), alr5065 (murC), all4316 (mraY), alr0093 (hcwA, amiC2) or all3826 (a penicillin binding protein, PBP). Several of the predicted targets corresponded to proteins known to be located in the outer membrane (21), including alr0834, encoding a homolog of the two previously validated targets of Yfr1 in Prochlorococcus (24), alr4550 and all4499 (OprB-like porins), all2158 and all4026 (TonB dependent transporters), proteins related to the biogenesis of outer membrane (alr2269,

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Omp85) or a component of a TolC-like secretion system (alr2887). In addition, the proteins encoded by all0089 and all3310, also predicted targets, have been previously described as located in the outer membrane although their possible function is unknown (31). Among the top 50 predicted targets were also two genes (patN and conR) which are related to cell wall maintenance and also involved in certain aspects of heterocyst differentiation. Biased inheritance of PatN, located in the cytoplasmic membrane, has been related to the differentiation of certain cells into heterocysts (32), whereas ConR is essential for proper septum formation between cells (33).

Figure 1. Verification of the interaction between Yfr1 and several mRNAs using an in vivo reporter system in E. coli. (A) Different versions of Yfr1 used to validate Yfr1 interaction with several targets. The top line is the wild type sequence. Mutations introduced in Yfr1 at positions 31-32 (AC to UG, Yfr1_UG) and 27-30 (CCUC to AAAA, Yfr1_AAAA) are shown in red and purple, respectively. The fully conserved motif of Yfr1 is shown shaded in orange. (B) Fluorescence of E. coli cultures bearing different combinations of plasmids expressing wild-type or mutated versions of Yfr1 or a control RNA (pJV300) and different 5’-UTRs fused to sfgfp. The data are presented as the mean and standard deviation of the results from 8 independent colonies after subtraction of fluorescence in cells bearing pXG0 and normalized for cell density (A600). Results are presented in two graphs according to the different scale required.

We have verified the interaction between Yfr1 and 11 selected targets (shown in grey in Table S1), using a heterologous reporter system (34), in which the 5’-UTR of the predicted

target mRNAs (plus sequences encoding the first 10-20 amino acids of the corresponding protein) is translationally fused to the gene sfgfp and co-expressed in E. coli with Yfr1 or with a

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control, unrelated RNA. We were able to measure fluorescence of cells carrying fusions to sfGFP of every target, indicating that the translation initiation regions of these mRNAs were functional in E. coli. In all cases analyzed, except for alr0093::sfgfp, the fluorescence of cells carrying sfgfp fusions significantly decreased when Yfr1 was co-expressed, indicating a negative effect of Yfr1 on expression of sfGFP (Fig. 1B). In contrast, in the case of alr0093::sfgfp, the fluorescence was higher in the presence of Yfr1.

The interactions between Yfr1 and the 5’UTRs of the eleven target mRNAs analyzed, as predicted by IntaRNA (35), are shown in Figure S2. In all cases the predicted interaction involves the conserved region of Yfr1 and takes place in a region of the mRNA located just upstream of the translational start of the corresponding gene except for alr2269 and alr0093, where the interaction is predicted further upstream. alr0093 is the only fusion whose expression was activated by interaction with Yfr1 (Fig. 1B).

To verify that the interactions with the predicted targets involved the conserved region of Yfr1 we designed two mutated versions of Yfr1, one altered in positions 31-32 (AC to UG, Yfr1_UG) and another one altered in positions 27-30 (CCUC to AAAA, Yfr1_AAAA) (Fig. 1A). In all cases Yfr1_UG had a slightly weaker effect than wild type Yfr1 in the level of the GFP fluorescence of strains bearing the sfGFP fusions. In contrast, mutations introduced in Yfr1_AAAA strongly affected the magnitude of the change in GFP fluorescence of the fusion proteins, when compared with the change due to coexpression with wild type Yfr1 (Fig. 1B). These results confirmed that the highly conserved region of Yfr1 was involved in the interaction between Yfr1 and its targets, as previously reported for Yfr1 in Prochlorococcus (24).

We further analyzed the interaction between Yfr1 and the 5’-UTR of two genes involved in peptidoglycan synthesis, all4316 (mraY), and alr5065 (murC). We designed compensatory mutations in the 5’-UTRs of the two genes that would restore the interaction with Yfr1_UG (Fig. 2A-B). When Yfr1_UG was combined with the mutated versions of the 5’-UTRs containing the designed compensatory changes, base pairing was restored resulting in a stronger change of fluorescence with respect to cells co-expressing the unrelated control RNA than when Yfr1_UG was combined with the wild type version of the 5’-UTRs (Fig. 2C-D).

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Figure 2. Interaction of Yfr1 and the 5’-UTRs of alr5065 and all4316 in E. coli. (A-B) Interaction predicted by IntaRNA between Yfr1 and the 5’-UTR of all4316 (A) and alr5065 (B). Nucleotides of 5’-UTRs are numbered with respect to the start of the coding sequence (start codons are indicated in bold and underlined). Mutations introduced in Yfr1 at positions 31-32 (AC to UG, Yfr1_UG) and compensatory mutations introduced in the 5’-UTRs of the mRNAs are shown in red and blue, respectively. The additional mutations introduced in Yfr1 at positions 27-30 (CCUC to AAAA, Yfr1_AAAA) are shown in purple. (C-D) Fluorescence of E. coli cells bearing combinations of plasmids expressing different versions of Yfr1 and the wild type or mutated versions of the 5’-UTR of all4316 (C) or alr5065 (D). The data are presented as mean and standard deviation of the results from 8 independent colonies after subtraction of fluorescence in cells bearing pXG0 and normalized for cell density (A600).

Yfr1 affects all4316 (mraY) and alr5065 (murC) expression in Nostoc sp. PCC 7120.

In order to study the function of Yfr1 in Nostoc, we prepared strains with altered levels of Yfr1 (Table S2). Because Yfr1 accumulates at relatively high levels in the wild type strain, to overexpress Yfr1 we introduced in Nostoc a plasmid designed for very strong expression from the trc promoter (strain OE_Yfr1) (Fig. 3A). In this plasmid, the segment cloned downstream of the trc promoter is transcribed until the T1 transcriptional terminator from the rrnB gene of E. coli (36). A six-nucleotides tag was introduced between the transcriptional start site of the trc promoter and the DNA segment encoding Yfr1 so that the native endogenous molecules of Yfr1 could be distinguished from Yfr1 molecules expressed from the trc promoter based on their length. In order to reduce the amount of Yfr1 we followed a strategy used previously (37), and transformed Nostoc with a plasmid bearing the Yfr1 sequence in reverse orientation

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downstream the trc promoter, so that its transcription generates a perfect antisense to Yfr1 (Fig. 3A) that would act as a sponge neutralizing Yfr1 (strain OE_as_Yfr1). As a control we used a Nostoc strain with a plasmid without insert between the trc promoter and the terminator (OE_C).

Figure 3. Expression of Yfr1, all4316 and alr5065 in OE_C, OE_Yfr1 and OE_as_Yfr1 Nostoc strains. (A) Scheme of plasmids pMBA51 (OE_C), pMBA48 (OE_Yfr1) and pMBA49 (OE_as_Yfr1). The transcription start site (bent arrows), T1 terminator (large stem loops), Rho-independent terminator of Yfr1 (small stem loops), trc promoter, probe for Yfr1 (red thick line) and sequences corresponding to Yfr1 (red arrows) are indicated. (B, C, E) Northern blots with RNA from three independent clones of OE_C, OE_Yfr1 and OE_as_Yfr1 strains grown in the presence of ammonia and hybridized with probes for Yfr1 (B), all4316 (C), alr5065 (E) and 5S RNA (B) or rnpB (C, E) as loading controls. Endogenous Yfr1 (black triangle) and Yfr1 expressed constitutively from the trc promoter (red triangle) are indicated. (D, F) Quantification of all4316 (D) or alr5065 (F) expression. Data are presented as the mean ± standard deviation of the signal in the largest intact band normalized to the rnpB signal (three individual clones of each strain). (*) p<0,05; (**) p<0,01, t-test.

We analyzed the accumulation of Yfr1 in Nostoc strains bearing the above described constructs by northern blot hybridization, using three independent clones of the OE_C, OE_Yfr1 and OE_as_Yfr1 strains grown in the presence of combined nitrogen (ammonium). Expression

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of Yfr1 under the trc promoter clearly exceeded endogenous Yfr1 expression whereas transcription of the sequence antisense to Yfr1 led to complete depletion of endogenous Yfr1 (Fig. 3B). In the OE_Yfr1 strain, termination of the strong transcription from the trc promoter at the Rho-independent terminator of Yfr1 was only partial and most transcripts were terminated at the T1 transcriptional terminator (56 nucleotides downstream) and appeared as longer molecules in northern blots (Fig. 3B).

We then tested the accumulation of the mRNAs of all4316 and alr5065 in the OE-C (control), OE_Yfr1 and OE_as_Yfr1 strains. The accumulation of intact all4316 and alr5065 mRNAs was significantly stronger in the OE_as_Yfr1 strain (depleted of Yfr1) than in the OE_C strain (Fig. 3C-F). There was also a slightly reduced accumulation of intact alr5065 mRNA in OE_Yfr1 strain with respect to the control strain (Fig. 3E-F), and degradation products of all4316 were clearly observed in strain OE_Yfr1. Taken together, these results indicated that Yfr1 negatively affects the accumulation of all4316 and alr5065 mRNAs in Nostoc sp. PCC 7120. This is the expected result if inhibition of their translation by Yfr1 results in indirect destabilization of the mRNAs, in agreement with the results obtained in E. coli (Figs. 1 and S3).

Strains with altered levels of Yfr1 show altered cell wall integrity and peptidoglycan synthesis.

We have validated the interaction between Yfr1 and the 5’-UTRs of several mRNAs encoding proteins related to cell wall (transporters and proteins located in the outer membrane) and peptidoglycan biosynthesis or remodeling. In order to verify the physiological relevance of Yfr1, we tested the effects of several harmful compounds that could affect the growth of strains with a compromised cell wall on the above-described strains with altered levels of Yfr1. In contrast to the OE-C strain, neither the OE_Yfr1 nor the OE_as_Yfr1 (Yfr1 depleted) were able to grow on plates containing 100 ng/mL vancomycin, an antibiotic that binds to the nascent peptidoglycan chains (Fig. 4A). In addition, OE_Yfr1 grew slightly worse than the control strain in plates containing SDS or erythromycin (Fig. 4A).

We also visualized the septa between cells by incorporation of a fluorescent derivative of vancomycin (Van-FL), that binds to nascent peptidoglycan chains (38). In comparison to the control strain OE-C, that only showed fluorescent septa between individual cells, OE_Yfr1 also showed fluorescent septa in the middle of cells that had not completed division (Fig. 4B). Most of the septa in the OE-Yfr1 strain were wider than those in the OE_C strain, suggesting cell division was not properly completed. In contrast, strain OE_as_Yfr1 showed very narrow septa between cells that had completed division suggesting a faster completion of the septa.

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Figure 4. Functionality of cell wall in strains with altered levels of Yfr1. (A) Growth of OE_C, OE_Yfr1 and OE_as_Yfr1 strains in media containing nitrate (BG11) and supplemented with the indicated substances. Pictures were taken after 10 days of incubation at 30ºC. (B) Fluorescence microscopy images of Van-FL stained filaments of OE_C (left), OE_Yfr1 (center) and OE_as_Yfr1 (right) strains.

Yfr1 may affect heterocyst differentiation.

The differentiation of heterocysts involves important morphological changes, including the secretion of specific components of envelopes (HEP and HGL) outside the outer membrane. Peptidoglycan remodeling seems essential for the correct deposition of these

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envelopes as well as for the proper communication between vegetative cells and heterocysts (10-12, 14, 15). In order to analyze completion of heterocyst differentiation in strains OE_Yfr1 and OE_as_Yfr1, we tested their growth in plates containing different nitrogen sources (Fig. 5A). Whereas strain OE_as_Yfr1 showed no difference with respect to the OE_C strain, strain OE_Yfr1 grew worse than the control strain in plates without combined nitrogen, a nutritional condition that requires differentiation of functional nitrogen-fixing heterocysts. Strain OE_Yfr1 was unable to grow in liquid media without combined nitrogen (data not shown). The visualization of filaments of strain OE_Yfr1 streaked on top of plates of media lacking combined nitrogen showed patterned differentiation of heterocysts but the filaments appeared broken between heterocysts and adjacent vegetative cells (Fig. 5B).

Figure 5. Growth of strains with altered levels of Yfr1. (A) Growth of OE_C, OE_Yfr1 and OE_as_Yfr1 strains in media - + lacking nitrogen (N2), or containing nitrate (NO3 ) or ammonium (NH4 ). Pictures were taken after 10 days of incubation at 30ºC. (B) Bright field images of filaments from the OE_C strain and from two independent clones of strain OE_Yfr1 streaked on top of BG110 plates. The broken connections between heterocysts and vegetative cells are indicated with black triangles.

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DISCUSSION

Yfr1 is an sRNA conserved in all cyanobacterial genomes (27, 28). The conservation of an sRNA in organisms with such a wide variety of morphologies, ecological niches and developmental processes, may suggest a regulatory function of a general aspect of the physiology of cyanobacteria. The prediction of targets by CopraRNA (29, 30) for Yfr1 homologs in heterocyst-forming cyanobacteria showed a remarkable enrichment in transporters and proteins located in the outer membrane as well as proteins involved in the synthesis or remodeling of peptidoglycan (Table S1). Target prediction was performed using genomes of heterocystous strains and in fact, among the TOP 50 predicted targets there are some genes of unknown function, such as alr4714, asl4743, all0997 and alr0255, that appear conserved only in this cyanobacterial clade (39).

Using a heterologous assay in E. coli we have demonstrated the effect of Yfr1 on expression of eleven selected predicted targets and have verified that Yfr1 mostly downregulates the expression of the targets (ten cases), but it can also exert a positive regulation, as in the case of alr0093 (Fig. 1B). We could demonstrate that regulation was exerted via base-pairing of the highly conserved motif of Yfr1 with the mRNAs (Fig. 1 and 2). Most of the predicted interaction sites were located in the translation initiation region (Fig. S2). However, the interaction between Yfr1 and alr0093 was located far upstream from the start codon (Fig. S2), opening the possibility that the positive regulation of this particular mRNA is operated through a conformational change in its 5’-UTR that might improve translation. Using compensatory mutations, we have mapped the interactions between Yfr1 and the 5’-UTR of all4316 and alr5065 to the ribosome binding region (Fig. 2). These interactions are consistent with a negative regulation based on interference with ribosome access. In fact, the reduction in the corresponding RNA levels is only around 30-50% (Fig. S3A-B) whereas the fluorescence of translational reporters fused to the 5’ UTRs of all4316 and alr5065 is reduced more than 90% in the presence of Yfr1 (Fig. S3C), further suggesting a mechanism involving translational interference rather than alteration of mRNA stability.

In order to assess the effects of Yfr1 in Nostoc, we overexpressed Yfr1 and anti-Yfr1 RNA under the strong and constitutive trc promoter (Fig. 3). Indeed, by analyzing Nostoc strains with altered levels of Yfr1 we observed a higher accumulation of full-length mRNAs for all4316 and alr5065 in strain OE_as_Yfr1 (Fig. 3C-F), that is depleted of Yfr1 (Fig. 3B), than in the control OE-C strain. This result was consistent with the negative regulation exerted by Yfr1 on these targets as validated in the E. coli system (Fig. 2).

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Vancomycin binds to the D-alanyl-D-alanine terminus of nascent glycan chains preventing the cross-linking between two glycan strands (40). Strains OE_Yfr1 and OE_as_Yfr1 were more sensitive to vancomycin than strain OE_C strain (Fig. 4A), suggesting that both strains have alterations in peptidoglycan integrity, consistent with altered amounts of murC and mraY mRNAs. In addition, the poor growth of the OE_Yfr1 strain in plates containing SDS (a detergent) or erythromycin (a macrolide antibiotic) suggested additional defects in the integrity and permeability of the envelopes of this strain. This result is consistent with a negative regulation of Yfr1 on alr2269 mRNA (Omp85), since an alr2269 mutant showed greater sensitivity to SDS and erythromycin (19). When we measured the incorporation of Van-FL, a fluorescent derivative of vancomycin, we observed opposite phenotypes for the OE_Yfr1 and OE_as_Yfr1 strains (Fig. 4B). OE_Yfr1 had wider, frequent nascent septa in the middle of cells that had not finished its previous division whereas OE_as_Yfr1 had narrower septa and only some new septa were starting its completion. These results might be consistent with altered peptidoglycan synthesis and remodeling mediated by a negative regulation by Yfr1 of all4316 (murC) and alr5065 (mraY) and a positive regulation of alr0093 (amiC2). Modulation of the amounts of the corresponding enzymes could facilitate a faster completion of the peptidoglycan layer in OE_as_Yfr1 and a slower completion of the peptidoglycan layer in OE_Yfr1 strain, although these strains could differ also in other aspects of cell division. In fact, ConR, a protein of the LytR-CpsA-Psr superfamily involved in septum formation (33), is one of the targets of Yfr1 verified in this work (Fig. 1).

Heterocyst differentiation involves important morphological changes in which synthesis and remodeling of peptidoglycan play an important role. Two of the validated targets of Yfr1 (amiC2 and murC) are necessary for proper diazotrophic growth (11-13). In addition, mraY (also among the targets of Yfr1 validated in this study) has a complex promoter region in which one transcription start site is specifically upregulated in heterocysts (41), suggesting a role of this enzyme in heterocyst differentiation. Strain OE_Yfr1 was unable to grow in liquid medium without combined nitrogen and grew very poorly on solid medium without combined nitrogen (Fig. 5A). Filaments of OE_Yfr1 plated on top of medium without combined nitrogen showed that although the strain was able to differentiate heterocysts with a normal pattern, the heterocyst-vegetative cells connections appeared frequently disrupted (Fig. 5B). These results suggest that the regulation exerted by Yfr1 at the level of a general aspect of the physiology of Nostoc sp. PCC 7120 (bacterial envelopes), is also critical for the differentiation of functional mature heterocyst.

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Finally, expression of most previously studied cyanobacterial sRNAs, such as NsiR1 (42, 43), NsiR4 (44), IsaR1 (45), PsrR1 (46) or NsrR1 (47) transiently changes in response to certain environmental conditions, including light or availability of different nutrients. Expression of Yfr1 however is relatively high in Nostoc sp. PCC 7120 under standard laboratory conditions and its expression did not significantly change in response to high light or nitrogen availability, the two conditions we tested (Fig. S1). In Synechococcus elongatus PCC 6301, Yfr1 accumulates up to 18,000 copies per cell and its expression barely changes under different stresses (26). Yfr1 expression only slightly changes under tested stress conditions in Prochlorococcus MED4 (25). Therefore, the question arises how the regulation exerted by Yfr1 on its targets is modulated. One possibility is that the regulatory effects of Yfr1 depend on its controlled sequestration by an RNA-binding protein or another sRNA that could act as a trap. In the unicellular cyanobacterium Prochlorococcus, the regulation of Yfr1 occurs through sequestration by another conserved sRNA, Yfr2, that contains a conserved region partially complementary to Yfr1 and whose accumulation was found to respond to changes in the nitrogen availability (25). In Nostoc, Yfr2 is also upregulated under nitrogen deprivation (16) and we have carried out EMSA assays showing that Nostoc Yfr2 also interacts with Yfr1 in a similar way to that described in Prochlorococcus (Fig. S4). Thus, under nitrogen deprivation or other stresses Yfr2 transcribed from one or several of the four repeats found in the genome of Nostoc sp. PCC7120 could bind to the conserved region of Yfr1 preventing the interaction of Yfr1 with its target mRNAs (Fig. 6). Overexpression of Yfr1 in strain OE_Yfr1 might be buffered by interaction with Yfr2, therefore leading to changes that are less evident than those observed in the E. coli system, in which Yfr2 is absent, while Yfr1 is expressed from a high copy number plasmid.

The regulatory model proposed here implies that under nitrogen stress Yfr2 would reduce the inhibitory effect of Yfr1 on a number of proteins required for cell wall changes occurring during heterocyst development. It is possible that additional regulatory mechanisms ensure specific enrichment of those proteins in heterocysts, as discussed above for mraY. The results presented here shed light on a general regulatory network that assures that proper amounts of different proteins related to cell wall biosynthesis and remodeling are present in Nostoc cells under different circumstances, and specifically during heterocyst development.

The use of regulatory RNAs to control the amounts of outer membrane proteins is well described in Enterobacteria (48, 49). Enzymes involved in peptidoglycan biosynthesis have also been found to be controlled by a small RNA (50). Our work shows that also in Cyanobacteria similar mechanisms operate for the coordinated regulation of outer membrane proteins and peptidoglycan biosynthesis enzymes through Yfr1.

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Finally, though Yfr1 is universally conserved in cyanobacteria, it appears that different regulatory functions are ascribed to Yfr1 in evolutionary distant strains. While in Prochloroccocus Yfr1 regulates genes involved in carbon and nitrogen metabolism (25), here we show that in a heterocyst-forming strain Yfr1 regulates genes related to cell wall synthesis and remodeling. Because most unicellular cyanobacteria seem to have evolved from non-heterocystous filamentous strains (51) it would be interesting to analyze the targetome of Yfr1 in non- heterocystous filamentous strains.

Figure 6. Yfr1 regulatory network. The sRNAs Yfr1 and Yfr2 (orange ovals) and the proteins encoded by Yfr1 target mRNAs (blue ovals) are shown within a schematic drawing of a cell (the thick light gray or dark gray lines indicate cytoplasmic and outer membrane, respectively). Proteins are located in different areas depending on its function or the physiological processes in which they may be involved; top right, peptidoglycan biosynthesis or remodeling, bottom right, transporters, bottom left, proteins related to heterocyst differentiation and top left, other functions. Positive regulation is indicated by a blue arrow, negative regulation by black lines with blunt ends. The hypothetical negative regulation of Yfr1 by Yfr2 is indicated by a red line with blunt end and a red question mark. Possible regulation of Yfr2 by environmental signals is indicated with a dashed arrow. OM: outer membrane; CM: cytoplasmic membrane.

MATERIAL AND METHODS

Strains and growth conditions.

Nostoc sp. PCC 7120 wild type and OE_C, OE_Yfr1 and OE_as_Yfr1 strains (Table S2)

were grown photoautotrophically at 30ºC in BG11 medium (52) lacking NaNO3 but containing

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3.5 mM NH4Cl and 7 mM N-[Tris(hydroxymethyl)methyl]-2-aminoethanesulfonic acid-NaOH buffer (pH 7.5). For Northern blot analysis of Yfr1 expression under different conditions, cultures of Nostoc sp. PCC 7120 were bubbled with an air/CO2 mixture (1% v/v) and grown photoautotrophically at 30ºC in BG11 medium (52) supplemented with 10 mM NaHCO3 (BG11C) lacking NaNO3 but containing 6 mM NH4Cl and 12 mM N-[Tris(hydroxymethyl)methyl]-2-

+ aminoethanesulfonic acid-NaOH buffer (pH 7.5) (BG11C + NH4 ). To induce nitrogen deficiency, filaments were collected by filtration, washed and resuspended in nitrogen-free BG11 medium containing 10 mM NaHCO3 (BG110C). High light stress (HL stress) was induced by increasing light intensity from 50 µE m-2 s-1 to 500 µE m-2 s-1.

To test growth of strains with altered levels of Yfr1 under different conditions, liquid cultures of these strains growing in BG11 media were diluted to A750 0.17 and 10 µL of serial 5- fold dilutions were spotted on plates containing different nitrogen sources and/or different harmful compounds.

OE_C, OE_Yfr1 and OE_as_Yfr1 strains were grown in the presence of appropriate antibiotics at the following concentrations: streptomycin (Sm) and spectinomycin (Sp), 2 µg/mL each (liquid medium) or 5 µg/mL each (solid medium).

E. coli strains (Table S2) were grown in LB medium, supplemented with appropriate antibiotics (53).

Generation of Nostoc strains with altered levels of Yfr1.

Plasmids and oligonucleotides used in this work are described in Table S3 and Table S4, respectively.

We have used pMBA37 (36) as a vector for overexpression of Yfr1 or an antisense to Yfr1 (as_Yfr1). pMBA37 contains the trc promoter and the T1 terminator of the rrnB gene of E. coli as transcriptional terminator and allows the overexpression of a cloned sequence flanked by NsiI-XhoI sites. Sequences corresponding to Yfr1 and as_Yfr1 (Yfr1 in reverse orientation) were amplified using genomic DNA as template and oligonucleotides 575 and 576 or 577 and 578, respectively. After digestion of the PCR products with NsiI and XhoI at the sites provided by the oligonucleotides, the fragments were cloned into NsiI-XhoI digested pMBA37, rendering pMBA48 (Yfr1) and pMBA49 (as_Yfr1). pMBA51 (a plasmid that overexpresses a control RNA corresponding only to the T1 terminator under the trc promoter) (36), pMBA48 and pMBA49

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were introduced in Nostoc sp. PCC 7120 by conjugation (54) generating strains OE_C (control), OE_Yfr1 and OE_as_Yfr1, respectively (Table S2).

RNA isolation, Northern blot analysis and primer extension assays.

Total RNA was isolated using hot phenol as described (55) with modifications (27). Northern blot detection of Yfr1 was performed using 10% urea-polyacrylamide gels as described (56) and 7.5 µg of total RNA. Northern blot hybridization for mRNAs (all4316 and alr5065) was performed using 1% agarose denaturing formaldehyde gels and 10 µg of total RNA. All RNA samples were then transferred to Hybond-N+ membrane (GE Healthcare) with 20XSSC buffer. Strand-specific 32P-labeled probes were prepared with Taq DNA polymerase using a PCR fragment as template and oligonucleotides specified in Table S4 in a reaction with α-32P-dCTP and one single oligonucleotide as primer (corresponding to the complementary strand of the sRNA or mRNA to be detected). PCR fragments used as templates for Yfr1, all4316, and alr5065 probes were amplified from genomic DNA using oligonucleotides pairs 368/369, 430/431, and 448/449, respectively. Hybridization to rnpB (57) or 5S rRNA was used as a loading and transfer control.

Fluorescent vancomycin conjugate staining.

Filaments from 1 mL of liquid cultures growing in BG11 medium for 5 days were pelleted, washed, resuspended in 50 µL of PBS buffer, mixed thoroughly and incubated with vancomycin-FL, (Van-FL; BODIPYTM FL Conjugate, Invitrogen) or uncoupled vancomycin at 1 µg/mL for 1 hour in the dark. After the incubation, unlinked Van-FL or vancomycin were removed by washing twice with PBS buffer. Fluorescence was analyzed using a Leica HCX PLAN-APO 63X 1.4 NA oil immersion objective attached to a Leica TCS SP2 confocal laser-scanning microscope. Van-FL was excited at 488 nm by an argon ion laser and the fluorescent emission was monitored in the range of 500 to 530 nm. Samples incubated with uncoupled vancomycin were used to set a threshold to measure the specific fluorescence of Van-FL.

Computational methods.

Sequences of homologs of Yfr1 were taken from (27). CopraRNA (29, 30) was used for the prediction of the targets of Yfr1, using the homologs in the genomes of Nostoc sp. PCC 7120, Nostoc sp. PCC 7524, Anabaena variabilis ATCC 29413, Nostoc sp. PCC 7107, Calothrix sp. PCC

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7507, Nostoc azollae 0708, Nostoc punctiforme PCC 73102, Calothrix sp. PCC 6303, Cylindrospermum stagnale PCC 7417 and Anabaena cylindrica PCC 7122. Prediction of the interaction site between Yfr1 and the 5’-UTR of several predicted targets in Nostoc sp. PCC 7120 was performed using IntaRNA (35). Alignment of Yfr2 homologs was made using Clustal Omega (58). Secondary structures of Yfr1 and Yfr2-1 and their interaction were predicted by RNAcofold (59).

Reporter assay for in vivo verification of targets.

We used the reporter assay described in (60) and fusions to the gene encoding superfolder GFP (sfGFP) in plasmids pXG10-SF or pXG30-SF (34) for experimental target verification in E.coli (Table S3). In this system both the GFP fusions and Yfr1 are transcribed constitutively.

5’-UTR of monocistronic targets were cloned in pXG10-SF from their corresponding TSS according to (16) to 30-60 nucleotides within the coding region. For targets that could be co- transcribed with a gene located upstream, the last 60 nucleotides of the upstream gene, together with the whole intergenic region plus 60 nucleotides within the coding region of the target gene were cloned in pXG30-SF. To facilitate translation in E. coli, GTG start codons were replaced by ATG using overlapping PCR and oligonucleotides specified in Table S4. PCR fragments containing the region to be cloned were amplified using genomic DNA as template and oligonucleotides specified in Table S4. Fragments were digested with NsiI and NheI and cloned into pXG10-SF or pXG30-SF treated with the same enzymes, resulting in translational fusions of the different targets to sfGFP (Table S5).

To express Yfr1 in E. coli, the sequence encoding Yfr1 was amplified from genomic DNA using primers 422 (5’ phosphorylated) and 423. The PCR product was digested with XbaI and fused to a plasmid backbone that was amplified from pZE12-luc with primers PLlacOB and PLlacOD (60) and digested with XbaI, rendering pMBA1 (Table S6).

For the mutagenesis of Yfr1 and the 5’-UTRs of all4316 and alr5065, mutations were introduced by overlapping PCR with primers containing the desired changes (Table S4) and the fragments were cloned in the same way as the corresponding wild type versions. The specific mutations were designed based on changes in the hybridization energies predicted by IntaRNA (35).

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Combinations of plasmids bearing fragments encoding Yfr1 (or its mutated versions) and the 5’ UTRs of target genes (or mutated versions) were introduced in E. coli DH5α. Plasmid pJV300 was used as a control expressing an unrelated RNA. Fluorescence measurements were done with a microplate reader (Varioskan) using liquid cultures from eight individual colonies of cells carrying each plasmid combination, as previously described (29).

In vitro transcription of RNA and EMSA essay.

RNAs were transcribed in vitro with MEGAscript High Yield Transcription Kit (AM1333, Ambion). The DNA templates for the transcription of Yfr1 and Yfr2 were generated by PCR with a primer that includes a T7 promoter sequence upstream the 5’-end of the corresponding RNA, and a primer matching the 3’-end of the RNA (Table S4). The template used for these PCR amplifications was genomic DNA. After in vitro transcription, RNAs were treated with DNase I and purified by sequential phenol, phenol/chloroform and chloroform extractions, ethanol- precipitated at −20 °C, and washed with 70% ethanol.

For EMSA, 200 ng of each in vitro transcribed Yfr1 and Yfr2 were combined in a volume of 5 µL, denatured for 1 min at 95 °C and chilled on ice for 5 min. After denaturing and chilling steps, 10x structure buffer (AM7118, Ambion) was added and the samples were incubated further for 15 min at 37 °C, before adding 1 µL of 50 % glycerol. Samples were run on 2.5 % agarose gel with TBE buffer 0.5 % at 50 V in a cold chamber.

ACKNOWLEDGEMENTS

We thank Iris Maldener and Rebeca Pérez (University of Tübingen, Germany) for the protocol for the fluorescent vancomycin assay, Claudia Steglich (University of Freiburg, Germany) for making available to us the information on the interaction between Yfr1 and Yfr2 prior to publication, and Claudia Steglich and Wolfgang Hess (University of Freiburg, Germany) for valuable discussions. This work was supported by grants BFU2013-48282-C2-1 from Ministerio de Economía y Competitividad, and BFU2016-74943-C2-1-P from Agencia Estatal de Investigación (AEI), Ministerio de Economía, Industria y Competitividad, both cofinanced by Fondo Europeo de Desarrollo Regional (FEDER), to AMMP. MBA is the recipient of a predoctoral contract from Ministerio de Educación, Cultura y Deporte, Spain (FPU014/05123 and EST16- 00088).

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SUPPLEMENTAL MATERIAL

Figure S1. Expression of Yfr1 in Nostoc sp. PCC 7120. (A) Northern blot analysis with total RNA extracted at different times after removal of combined nitrogen. (B) Northern blot analysis with total RNA from cells growing in the presence of ammonia at 50 µE m-2 s-1 and incubated at 500 µE m-2 s-1 (high light stress, HL) for the indicated times. All membranes were hybridized with probes for Yfr1 and 5S RNA as loading control.

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Figure S2. Interactions predicted by IntaRNA between Yfr1 and several mRNAs. Nucleotides of 5’UTRs are numbered with respect to the start of the coding sequence (start codons are indicated in bold and underlined). Positions of the mutations introduced in Yfr1_UG are shown in red. Positions of the mutations introduced in Yfr1_AAAA are shown in purple.

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Figure S3. Comparison of the effects of Yfr1 on expression of all4316 and alr5065 at the mRNA (A, B) and protein level (C) in E. coli. (A) Northern blot with RNA isolated from E. coli cells bearing plasmid pMBA4 (all4316-sfgfp) or pMBA7 (alr5065-sfgfp) and a plasmid expressing Yfr1 (pMBA1) or a control RNA (pJV300) hybridized with probes for all4316, alr5065, Yfr1 and 5S RNA as loading control. (B) Quantification of signals in Northern blots. Data are presented as the mean +/- standard deviation of the signal normalized to the 5S signal (three individual experiments). (C) Fluorescence of the E. coli cultures analyzed in (A-B). The data are presented as mean +/- standard deviation of the results from three cultures after subtraction of fluorescence in cells bearing pXG0 and normalized for cell density (A600). (*) p<0,05: (**) p<0,0001, t-test.

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Figure S4. Interaction between Yfr1 and Yfr2. (A) Clustal Omega alignment of the four Yfr2 RNAs encoded in the genome of Nostoc sp. PCC 7120. The region predicted to interact with Yfr1 is shaded in orange. (B) Secondary structures of Yfr1 (black) and Yfr2-1 (red) and their interaction as predicted by RNAcofold. (C) EMSA showing the interaction between in vitro transcribed Yfr1 and Yfr2. Equimolar amounts of Yfr1 and Yfr2 were combined and subjected to electrophoresis as described in the Material and Methods.

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Table S1. Top 50 predicted targets of Yfr1.

Gene Annotationa fdr p-value alr2458 alanine racemase 7,52E-09 2,14E-12 alr4550 porin-like, OprB-like 1,51E-08 8,60E-12 alr5065 (murC) UDP-N-acetylmuramate-L-alanine ligase 2,82E-08 2,40E-11 all4829 undecaprenyl-phosphate 2,64E-07 3,00E-10 galactosephosphotransferase all2158 ferrichrome-iron receptor (TonB-dependent 3,29E-07 4,68E-10 transporter) all4316 (mraY) phospho-N-acetylmuramoyl-pentapeptide 5,12E-07 8,72E-10 transferase alr2269 chloroplastic outer envelope membrane protein 6,58E-07 1,31E-09 homolog, Omp85, Toc75 all5094 oxidoreductase 7,30E-07 1,66E-09 all1835 unknown protein 2,39E-06 6,11E-09 alr0834 porin, ortholog of PMM1119 and PMM1121 3,05E-06 8,65E-09 all4499 porin-like, OprB-like 4,93E-06 1,54E-08 all2352 WD-40 repeat protein. 6,88E-06 2,35E-08 alr4077 hypothetical protein 7,09E-06 2,62E-08 all0187 (conR) LytR-CpsA-Psr superfamily 9,05E-06 3,68E-08 all1587 hypothetical protein 9,05E-06 3,86E-08 all0481 unknown protein 9,32E-06 4,24E-08 alr0068 adenylate kinase 1,37E-05 6,85E-08 all4220 hypothetical protein 1,37E-05 6,99E-08 alr3380 dolichol-phosphate mannosyltransferase 3,32E-05 1,81E-07 alr3654 uncharacterized RNA methyltransferase 3,32E-05 1,89E-07 alr2887 (hgdD) outer membrane channel, TolC-like, 4,54E-05 2,78E-07 alr2057 (aroE) shikimate 5-dehydrogenase 4,54E-05 2,96E-07 alr1805 hypothetical protein 4,54E-05 2,97E-07 all3826 peptidoglycan-binding protein 4,92E-05 3,36E-07 alr4812 (patN) heterocyst differentiation related protein 5,49E-05 3,90E-07 all4026 similar to TonB-dependent receptor 7,32E-05 5,41E-07 asr3279 unknown protein 8,67E-05 6,65E-07 alr0093 (amiC2, N-acetylmuramoyl-L-alanine amidase 9,51E-05 7,57E-07 hcwA) alr5049 unknown protein, putative OmpA/MotB system 9,87E-05 8,13E-07 all3669 unknown protein 1,05E-04 8,97E-07 alr4123 (prk) phosphoribulokinase 1,40E-04 1,23E-06 all1340 unknown protein 1,44E-04 1,31E-06 alr5030 hypothetical protein 1,99E-04 1,86E-06 all3902 hypothetical protein 2,35E-04 2,27E-06 all2116 hypothetical protein 3,54E-04 3,58E-06 alr4602 type I site-specific deoxyribonuclease chain S 3,54E-04 3,62E-06 alr2479 LytR-CpsA-Psr superfamily 3,80E-04 4,00E-06 alr4714 unknown protein, specific to heterocyst clade 3,85E-04 4,15E-06 alr4269 hypothetical protein 3,91E-04 4,33E-06

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alr3004 unknown protein 4,43E-04 5,03E-06 all0089 hypothetical protein 4,59E-04 5,34E-06 all0571 cyanophycinase 4,77E-04 5,69E-06 all0211 hypothetical protein 5,09E-04 6,21E-06 asl4743 unknown protein, specific to heterocyst clade 5,36E-04 6,70E-06 alr3955 thioredoxin 6,42E-04 8,21E-06 all3310 hypothetical outer membrane receptor for ferritin, 6,66E-04 8,71E-06 (TonB-dependent transporter) all0997 unknown protein, specific to heterocyst clade 7,50E-04 1,00E-05 all4106 probable oxidoreductase 7,91E-04 1,08E-05 alr0255 unknown protein, specific to heterocyst clade 7,98E-04 1,11E-05

aAnnotation based on NCBI (https://www.ncbi.nlm.nih.gov/), Cyanobase (http://genome.microbedb.jp/cyanobase/), CopraRNA results (1, 2) and previous studies.

Interactions verified in this work are highlighted in gray.

Table S2. Strains

Strain Description Reference Escherichia coli DH5α Used for routine (3) transformation

Nostoc sp. PCC 7120 Wild type Pasteur Culture Collection OE_C SmRSpR, pMBA51 inserted in (4) plasmid alpha. T1 terminator of E.coli rrnB gene expressed constitutively from trc promoter. OE_Yfr1 SmRSpR, pMBA48 inserted in This work plasmid alpha. Yfr1 expressed constitutively from trc promoter. OE_as_Yfr1 SmRSpR, pMBA49 inserted in This work plasmid alpha. Antisense to Yfr1 expressed constitutively from trc promoter.

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Table S3. Plasmids

Name Description Reference pMBA37 ApRSmRSpR, plasmid for the overexpression of transcripts from the trc (4) promoter and followed by the T1 terminator of E.coli rrnB gene, used as transcriptional terminator.

pMBA51 ApRSmRSpR, Control plasmid, expresses a 56 nt transcript derived from (4) the T1 terminator of E.coli rrnB gene from trc promoter. pMBA48 ApRSmRSpR, plasmid based on pMBA37 for the overexpression of Yfr1. This work pMBA49 ApRSmRSpR, plasmid based on pMBA37 for the overexpression of an This work antisense to Yfr1. pJV300 Control plasmid, expresses a ~50 nt transcript (5) derived from T1 terminator of E.coli rrnB gene. pXG0 CmR, control plasmid without sfgfp. (6) pXG10-SF CmR, plasmid for construction of translational (7) sfGFP fusions of monocistronic targets. pXG30-SF CmR, plasmid for construction of translational (7) sfGFP fusions of dicistronic targets. R pZE12-luc Ap , plasmid to express sRNAs under control of PLlac-O promoter. (8) pMBA1 ApR, plasmid based on pZE12-luc expressing Yfr1. This work pMBA13 ApR, same as pMBA1 but with AC to UG change at positions 31-32 of This work Yfr1 (mut 31_32). pMBA15 ApR, same as pMBA1 but with CCUC to AAAA change at positions 27-30 This work of Yfr1 (mut AAAA). pMBA2 CmR, sfGFP reporter plasmid based on pXG10-SF containing the all0187 This work 5’UTR plus sequences encoding the first 20 aminoacids (with mutation in start codon GTG -> ATG). pMBA3 CmR, sfGFP reporter plasmid based on pXG30-SF containing the all2158 This work 5’UTR plus sequences encoding the first 20 aminoacids. pMBA4 CmR, sfGFP reporter plasmid based on pXG30-SF containing the all4316 This work 5’UTR plus sequences encoding the first 20 aminoacids (with mutation in start codon GTG -> ATG). pMBA5 CmR, sfGFP reporter plasmid based on pXG10-SF containing the all4829 This work 5’UTR plus sequences encoding the first 17 aminoacids. pMBA6 CmR, sfGFP reporter plasmid based on pXG30-SF containing the alr0093 This work 5’UTR plus sequences encoding the first 20 aminoacids (with mutation in start codon GTG -> ATG). pMBA7 CmR, sfGFP reporter plasmid based on pXG10-SF containing the alr5065 This work 5’UTR plus sequences encoding the first 20 aminoacids. pMBA8 CmR, sfGFP reporter plasmid based on pXG10-SF containing the alr2458 This work 5’UTR plus sequences encoding the first 14 aminoacids. pMBA9 CmR, sfGFP reporter plasmid based on pXG10-SF containing the alr4550 This work 5’UTR plus sequences encoding the first 20 aminoacids. pMBA10 CmR, sfGFP reporter plasmid based on pXG10-SF containing the patN This work 5’UTR plus sequences encoding the first 20 aminoacids. pMBA11 CmR, sfGFP reporter plasmid based on pXG10-SF containing the alr0834 This work 5’UTR plus sequences encoding the first 20 aminoacids. pMBA17 CmR, same as pMBA7 but with a GT to CA change at position -21 and -22 This work of the 5'-UTR with respect to the start codon (alr5065 mut). pMBA19 CmR, same as pMBA4 but with a GT to CA change at position -22 and -23 This work of the 5'-UTR with respect to the start codon (all4316 mut). pMBA90 CmR, sfGFP reporter plasmid based on pXG10-SF containing the alr2269 This work 5’UTR plus sequences encoding the first 20 aminoacids (with mutation in start codon GTG -> ATG).

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Table S4. Oligonucleotides

Name Sequence (5’-3’) Used for 189 CGCACTGACCGAATTCATTAA Amplification of plasmid 190 GTGCTCAGTATCTTGTTATCCG backbone from pZE12-luc 356 AATCAGAACGCAGAAGCGGT Probe for 5S of E. coli 357 CGGCGGATTTGTCCTACTCA 368 AAAGGGAATAGGGAACGGCG Probe for Yfr1 369 CGGCTGCTGACATGCCTAAA 422 5’P-AGCGGAGACGCATGTTTCCGTTCAC Cloning of Yfr1 in pZE12-luc 423 GTTTTTTCTAGATACATTTGCAAAGAAGCTAAAGAATT 424 GTTTTATGCATAAACGGTAGAACATTTCAAGTG Cloning of all0187 5’-UTR with 425 GTTTTGCTAGCGAGATCCGGTATTTGTTGAGGC mutation in start codon (GTG- 426 TTAATTACAATTATCTCCCCACTCAACCA >ATG) in pXG10-SF 427 TAATTATGATTAAACAGGTTCAATGGTCA 428 GTTTTATGCATGCAGGTCACAAGACGGTTTT Cloning of all2158 5’-UTR in 429 GTTTTGCTAGCTCCACAAAGGCTGATGATAA pXG30-SF 430 GTTTTATGCATGACGACGATATCACCATAGA Cloning of all4316 5’-UTR with 431 GTTTTGCTAGCAGAAGCTAGACCAATACCAG mutation in start codon (GTG- 432 GCGTCCATAAAAGTTTCCCTTCACTCCAC >ATG) in pXG30-SF 433 CTTTTATGGACGCTAAATTATCGCCTA 434 GTTTTATGCATATGTGTGGTGTGGATTATTAAGG Cloning of all4829 5’-UTR in 435 GTTTTGCTAGCATCTTGCCGTAAGCGTCGTTT pXG10-SF 436 GTTTTATGCATCTTCAGTACCTCAAACGATAG Cloning of alr0093 5’-UTR with 437 GTTTTGCTAGCTGGTGACGATAGTAAGAAGA mutation in start codon (GTG- 438 AATTTCATAATCTTTTCTCCTGATGTGCG >ATG) in pXG30-SF 439 AGATTATGAAATTACACTGGTTACTATC 440 GTTTTATGCATATACTTTTGAAGTAAATATTACTTC Cloning of all4829 5’-UTR in 441 GTTTTGCTAGCATTTGTTTCTGTTATGGCAG pXG10-SF 442 GTTTTATGCATGTAATTGATCCTGTCGGTTA Cloning of alr2458 5’-UTR in 443 GTTTTGCTAGCTTGATGAGAAGCCATACTTG pXG10-SF 444 GTTTTATGCATATATGGGTACGTTTTTGCAT Cloning of alr4550 5’-UTR in 445 GTTTTGCTAGCTAGTGTTGCGCCTAAAAC pXG10-SF 446 GTTTTATGCATATCAGAATATTCATCTGTAAAAAA Cloning of patN 5’-UTR in 447 GTTTTGCTAGCCAAGAGAGCCGCCAGTCCAG pXG10-SF 448 GTTTTATGCATGTTATTTGTGATTGATGGTGTG Cloning of alr5065 5’-UTR in 449 GTTTTGCTAGCTATACCGCCGATTCCGATAA pXG10-SF 470 CTCCTCTGACCACACTCCGCCCGGACTA Mutagenesis of Yfr1 to 471 TGTGGTCAGAGGAGTGAACGGAAACATG generate Yfr1_UG 484 ATTAAACAGTGGAGTGAAGGGAAACTTT Mutagenesis of all4316 5’-UTR 485 CTCCACTGTTTAATAAAACTGAATTATA to generate all4316 mut 486 TGTGGTCAGGTGATAAGAGGAGTAATCA Mutagenesis of alr5065 5’-UTR 487 ATCACCTGACCACACCATCAATCACAAA to generate alr5065 mut 491 TCACTAAAAACACCACACTCCGCCCGGAC Mutagenesis of Yfr1 to 492 GGTGTTTTTAGTGAACGGAAACATGCGTC generate Yfr1_AAAA 575 GTTTTATGCATAGCGGAGACGCATGTTTCC Cloning of Yfr1 in pMBA37 576 GTTTTCTCGAGGGAACCGCCCGAACAGTAGT 577 GTTTTATGCATGGAACCGCCCGAACAGTAGT Cloning of as_Yfr1 in pMBA37 578 GTTTTCTCGAGAGCGGAGACGCATGTTTCCG 874 GTTTTATGCATACCTTTGGCATAAACAAGCAAT Cloning of alr2269 5’-UTR with 875 GTTTTGCTAGCAGTTTGGGCATTTGCAGTTAATGA mutation in start codon (GTG- 876 CAGCCGCCATTAATACGGGAGATAAACGCA >ATG) in pXG10-SF 877 TCCCGTATTAATGGCGGCTGTAGCAATCAC 903 TAATACGACTCACTATAGGGAGCGGAGACGCATGTTTCC G

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904 PCR of template for in vitro AAGGAACCGCCCGAACAGTAG transcription of Yfr1 905 TAATACGACTCACTATAGGGTGTGAGGAGCAAGTTGAAA PCR of template for in vitro 906 AAAAAGCATCTGGGAGTGTTGC transcription of Yfr2

Restriction sites used for cloning (underlined), point mutations introduced (red) and 5’ monophosphate (5’P) are indicated.

Table S5. Sequences of inserts in the sfGFP fusion plasmids.

Plasmid Sequence Description

pMBA2 atgcatAAACGGTAGAACATTTCAAGTGTACAGTGCGGTCTTACAC all0187 TGGCAAGATTGGTAGCTTGCTTTAACATAAATTTTAGGGAGTGGTT (mutation in GAGTGGGGAGATAATTATGATTAAACAGGTTCAATGGTCAGATAAT start codon) CAGGTCGCGCCTCAACAAATACCGGATCTCgctagc pMBA3 atgcatGCAGGTCACAAGACGGTTTTTGGATCATAACTGGCGGTTT all2158 TTGGATAGACACCCTGCGAGTAACTTCCCTATACTACTTTAACTGC TATTAAGAATTATTAGCAGTAATAGGCTGACTTTGATAAGTTTATT GTGTGGTGTGAGGAACGATGAAGCGTTGGTGTTTGTCCCCCAGTAT TCATTTATGGTTTATCATCAGCCTTTGTGGAgctagc pMBA4 atgcatGACGACGATATCACCATAGACGAAGATTAGTCTTTTGAGA all4316 CTGGGACTCTGTTATATATAGTCCCTATAATTCAGTTTTATTAAAG (mutation in TGTGGAGTGAAGGGAAACTTTTATGGACGCTAAATTATCGCCTAAT start codon) CAAGGATTAAATATTTCTGGTATTGGTCTAGCTTCTgctagc pMBA5 atgcatATGTGTGGTGTGGATTATTAAGGAGTATGATGACTGCCCA all4829 GAGTTCACTCCTCTCCGGCAAACGACGCTTACGGCAAGATgctagc pMBA6 atgcatCTTCAGTACCTCAAACGATAGATAATCAGTCAGTCTTCAC alr0093 AAACTTATAATCATCCTGATTGCCACACTATCCCTGTTATTCCTGA (mutation in AGATTTTGTGTGAATACTAACCACATCCTTATATTCTGAACCTGGA start codon) TCTTATGCCTTCACGACATGGCAAAATCAAAAGGGCGTGTTAAGAT GTGTCGCCATTAGTTGTATTTTTATACCAGCGCAAAATTTTTATTG CAGTAAAGCTGAAGTTTGAACGGTGATGGTGTGAGGATTAGCCAAC ATTTTGGAGCCTTAACGGCTCTCATGAACGCACATCAGGAGAAAAG ATTATGAAATTACACTGGTTACTATCGTCACCAgctagc pMBA7 atgcatGTTATTTGTGATTGATGGTGTGGTGTGGTGATAAGAGGAG alr5065 TAATCAATGAATAATAGTGTAGATTTTGGCGGTAGACCATTTCATT TTATCGGAATCGGCGGTATAgctagc pMBA8 atgcatGTAATTGATCCTGTCGGTTATTGGTAATGGCTCAGAACCC alr2458 CTATTAATTGGCTATTACCTCAGTCAAAAACAGAAGTTTAGCTCTT ATTACTGCTAGTTATTAGTAGCAAAAAGAACCTGTTAATTTTAATT AAGAGGTGGAGTATTGAGTAAAATACACTAGGTTCCAGTCTCTAAC CCCCTCCCTGATCAACATCTGAACGAGAAAATTTCAAAATTTTACT CAAGTATGTTGTATCTGTTGCGAAATGCGAGTTAAACTTCCTGATT AACTCTGATGTGGCTATAAAACCAATTCGCAATAGCCTGCGGCAAG GCTAACGCCAACGCAATTCGGAATAAATACGCATTTAAAAAGCAGT GGTGTGATAAGGGGGAGTGAAATGTTAAGTCGCCAACAAGCCTCAA GTATGGCTTCTCATCAAgctagc pMBA9 atgcatATATGGGTACGTTTTTGCATAAGTTAATTGCATTAATTTT alr4550 TAATACTAAGTTAATCAATCATCAGCTTGGCAAAAACTAAAATTGT AATAGTGGCAACTATGCCTCATCAAAAGATTTCAATAAGATGTCAC ATTTTTGAGAGTAGCTGCAACAAGAACAATTATCTATACAAGGTGT GAGGAGAAAAGTAAAAATGTCTAATCTATTGTGGAAATCCCTAGTG GTTAGCCCTGCTGTTTTAGGCGCAACACTAgctagc pMBA10 atgcatATCAGAATATTCATCTGTAAAAAAACTAGCTGAAACAGCA alr4812 GCTTAATATTTTGGGTATTATGTGAATTAAGTAGGCTAAAAGATTA

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TGAGTGTGAGAGTAGGTCAGTCAAACGTTATGGAATTTACATCGCA GATGAGGGGTGAAGGCGATGAGTGAAAGTATGGCATTTATCGGCGG TGTCGCCGTAGCTGGACTGGCGGCTCTCTTGgctagc pMBA11 atgcatATACTTTTGAAGTAAATATTACTTCAAATAAACTTGGTAA alr0834 ATCTATTGCCAAGAAATAAGATGTGTGTGAGTAAATTGTGAGGAGA AAAATGCAGAAAATATGGAAGTATTGGCTGGTTAACCCAGTAATTT GCAGCACAATGCTATTTTCTGGCGCTGCTGCCTTTGCAGGAGAAAC CCCTGCCATAACAGAAACAAATgctagc pMBA17 atgcatGTTATTTGTGATTGATGGTGTGGTCAGGTGATAAGAGGAG alr5065 mut TAATCAATGAATAATAGTGTAGATTTTGGCGGTAGACCATTTCATT TTATCGGAATCGGCGGTATAgctagc pMBA19 atgcatGACGACGATATCACCATAGACGAAGATTAGTCTTTTGAGA all4316 mut CTGGGACTCTGTTATATATAGTCCCTATAATTCAGTTTTATTAAAC AGTGGAGTGAAGGGAAACTTTTATGGACGCTAAATTATCGCCTAAT CAAGGATTAAATATTTCTGGTATTGGTCTAGCTTCTgctagc pMBA90 atgcatACCTTTGGCATAAACAAGCAATTGCCTGGTAATGGTGTGT alr2269 GGATGTGAAGAGGAAGAAAGGAATTAAAGACGATGCGTTTATCTCC (mutation in CGTATTAATGGCGGCTGTAGCAATCACAGCACCCTTGAGTAGTTCA start codon) TTAACTGCAAATGCCCAAACTgctagc

Nostoc sequences are capitalized, with black letters corresponding to 5’UTR and green letters corresponding to coding sequences, respectively. NsiI and NheI sites that were used for cloning are highlighted in magenta and yellow, respectively. Nucleotide changes with respect to wild type sequences are indicated in red.

Table S6. Sequences of inserts in plasmids used for expression of Yfr1 and its mutated versions.

Plasmid Sequence Description

pMBA1 AGCGGAGACGCATGTTTCCGTTCACTCCTCACACCACACTCCGCCCGGAC Yfr1 TACTGTTCGGGCGGTTCCTTATGTATTACATAAATTCTTTAGCTTCTTTG CAAATGTATCTAGA pMBA13 AGCGGAGACGCATGTTTCCGTTCACTCCTCTGACCACACTCCGCCCGGAC Yfr1_UG TACTGTTCGGGCGGTTCCTTATGTATTACATAAATTCTTTAGCTTCTTTG CAAATGTATCTAGA pMBA15 AGCGGAGACGCATGTTTCCGTTCACTAAAAACACCACACTCCGCCCGGAC Yfr1_AAAA TACTGTTCGGGCGGTTCCTTATGTATTACATAAATTCTTTAGCTTCTTTG CAAATGTATCTAGA

Grey shadowed letters indicate the Yfr1 sequence. Modified nucleotides are marked in red. XbaI restriction site used for cloning is shown in blue.

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SUPLEMENTARY REFERENCES

1. Wright PR, Georg J, Mann M, Sorescu DA, Richter AS, Lott S, Kleinkauf R, Hess WR, Backofen R. 2014. CopraRNA and IntaRNA: predicting small RNA targets, networks and interaction domains. Nucleic Acids Research 42:W119-W123. 2. Wright PR, Richter AS, Papenfort K, Mann M, Vogel J, Hess WR, Backofen R, Georg J. 2013. Comparative genomics boosts target prediction for bacterial small RNAs. Proc Natl Acad Sci U S A 110:E3487-3496. 3. Hanahan D. 1983. Studies on transformation of Escherichia coli with plasmids. J Mol Biol 166:557-580. 4. Olmedo-Verd E, Brenes-Álvarez M, Vioque A, Muro-Pastor AM. 2019. A heterocyst- specific antisense RNA contributes to metabolic reprogramming in Nostoc sp. PCC 7120. Plant Cell Physiol 60:1646-1655. 5. Sittka A, Pfeiffer V, Tedin K, Vogel J. 2007. The RNA chaperone Hfq is essential for the virulence of Salmonella typhimurium. Mol Microbiol 63:193-217. 6. Urban JH, Vogel J. 2009. A green fluorescent protein (GFP)-based plasmid system to study post-transcriptional control of gene expression in vivo. Methods Mol Biol 540:301-319. 7. Corcoran CP, Podkaminski D, Papenfort K, Urban JH, Hinton JC, Vogel J. 2012. Superfolder GFP reporters validate diverse new mRNA targets of the classic porin regulator, MicF RNA. Mol Microbiol 84:428-445. 8. Lutz R, Bujard H. 1997. Independent and tight regulation of transcriptional units in Escherichia coli via the LacR/O, the TetR/O and AraC/I1-I2 regulatory elements. Nucleic Acids Res 25:1203-1210.

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3.3 CHAPTER III A co-expression network to dissect the complex transcriptome of Nostoc sp. PCC 7120 during heterocyst differentiation

CHAPTER III. A co-expression network to dissect the complex transcriptome of Nostoc sp. PCC 7120 during heterocyst differentiation

Brenes-Álvarez, M., Mitschke, J., Olmedo-Verd, E., Georg, J., Hess, W.R., Vioque, A., and Muro- Pastor, A.M. (2019). Elements of the heterocyst-specific transcriptome unravelled by co- expression analysis in Nostoc sp. PCC 7120. Environmental Microbiology 21, 2544-2558. doi: 10.1111/1462-2920.14647.

3.4 CHAPTER IV NsiR1, a sRNA with multiple copies, regulates heterocyst differentiation

Classification: Biological Sciences, Microbiology

Title: NsiR1, a small RNA with multiple copies, regulates heterocyst differentiation in Nostoc sp. PCC 7120

Authors: Manuel Brenes-Álvarez, Marina Mingueta, Agustín Vioque1, and Alicia M. Muro-Pastor ORCID MBA: 0000-0002-6452-5557 ORCID AMMP: 0000-0003-2503-6336 ORCID AV: 0000-0002-3975-7348

Authors affiliation: Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and Universidad de Sevilla, Sevilla, Spain. aPresent address: Centro de Biología Molecular Severo Ochoa, CSIC-UAM, Nicolas Cabrera 1, 28049, Madrid, Spain.

1Corresponding Author: Agustín Vioque, Instituto de Bioquímica Vegetal y Fotosíntesis, Américo Vespucio 49, 41092, Sevilla, Spain Phone: +34-954489519; Email: [email protected] Keywords: cyanobacteria, differentiation commitment, Anabaena, HetR,

Author contributions: M.B.A., A.V., and A.M.M.P. designed research; M.B.A., M.M., A.V., and A.M.P. performed research; M.B.A., A.V., and A.M.P. analyzed data; and M.B.A., A.V., and A.M.P. wrote the paper.

The authors declare no conflict of interest.

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Abstract

Upon nitrogen starvation filamentous cyanobacteria develop heterocysts, specialized cells devoted to fixation of atmospheric nitrogen. Differentiation of heterocyst at semi-regular intervals requires complex structural and functional changes that are under control of the master transcriptional regulator HetR. NsiR1 (nitrogen stress-induced RNA 1) is a HetR- dependent non-coding RNA that is expressed in early steps of heterocyst development. In Nostoc sp. PCC 7120 there are twelve tandem copies of nsiR1 (nsiR1.1 to nsiR1.12), seven of them with identical sequence (nsiR1.3 to nsiR1.9) and the others slightly divergent. nsiR1.1 is transcribed antisense to the 5’ UTR of hetF, a gene required for heterocyst development. Here we show that binding of NsiR1.1 inhibits translation of the hetF mRNA by inducing structural changes in its 5’ UTR. Constitutive overexpression of NsiR1.1 or NsiR1.4 results in different phenotypes related to heterocyst development, suggesting that they regulate non-overlapping targets. We have identified gene alr3234, a hetP-like gene involved in regulation of commitment to heterocyst differentiation, as a target of NsiR1.4. A strain overexpressing NsiR1.4 commits to heterocyst development earlier than the wild type. The post-transcriptional regulation by NsiR1 of two genes related to the process of heterocyst differentiation and commitment, hetF and alr3234, adds a new level of complexity to the network of transcriptional regulation and protein- protein interactions involved in heterocyst differentiation.

Significance statement

Heterocysts are nitrogen-fixing specialized cells that appear at semi-regular intervals along cyanobacterial filaments upon nitrogen starvation. Heterocyst differentiation and patterning is a model system to study cell differentiation in multicellular prokaryotes. The differentiation process has a complex regulation, only partially understood, that includes transcriptional changes, factor diffusion between cells, and protein-protein interactions. This work adds to this complexity a non-coding RNA, NsiR1, that negatively regulates, at a post- transcriptional level, hetF and alr3234 (encoding a HetP-like protein), two genes that are part of the heterocyst differentiation regulatory network. Thus, regulation by a small RNA is a new element involved in the control of heterocyst differentiation.

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Introduction

Cyanobacteria are the only prokaryotes able to perform oxygenic photosynthesis and important contributors to global primary productivity. They constitute a coherent phylogenetic group with diverse morphology and wide ecological distribution. Nostoc sp. PCC 7120 (a.k.a Anabaena sp. PCC 7120) is a filamentous cyanobacterium that under nitrogen deprivation differentiates heterocysts, specialized cells devoted to fixation of atmospheric nitrogen. Heterocyst differentiation involves a complex developmental program (1-4) that is ultimately under control of NtcA, the global regulator of nitrogen assimilation (5), but also under control of HetR, a specific regulator of cell differentiation (6, 7). At the initiation of heterocyst differentiation, induction of the expression of NtcA and HetR depends on each other (8) and leads to increased expression of HetR in cells becoming heterocysts. Expression of hetR is regulated at several levels. hetR is transcribed from a complex promoter including several transcriptional start sites (TSSs), one of them specifically induced in cells becoming heterocysts (9, 10). HetR is a DNA-binding protein that positively regulates its own expression and the expression of many genes, including negative elements essential for the maintenance of the pattern of heterocysts along the filaments such as patS (11), hetN (12) and patX (13). The binding of the pentapeptide RGSGR, that is part of the sequence of PatS, blocks the DNA-binding activity of HetR (14) and signals its degradation (15). HetR is an autoproteolytic protein (16) that can also be degraded by the HetF protease (17, 18). A hetF mutant does not differentiate heterocysts, has an altered morphology with enlarged cells and accumulates a high amount of HetR (18). However, since hetF transcription seems unpatterned along the filaments (18), the question of how HetF may contribute to the patterned accumulation of active HetR remains open.

Because heterocysts are terminally differentiated non-dividing cells, a key aspect of differentiation is the so-called commitment point, a stage of differentiation from which the process is no longer halted by the addition of combined nitrogen. Several genes have been identified related to the regulation of commitment in Nostoc sp. PCC 7120, including hetP (19) and its homologs asl1930, alr2902 and alr3234 (20). Asl1930, Alr2902, and Alr3234 would interact with HetR and with each other, regulating commitment via protein-protein interactions (20). Mutant strains with genes alr3234 or asl1930 deleted reach the commitment point earlier than the wild-type strain, which suggests that Alr3234 and Asl1930 might function as negative regulators delaying commitment (20).

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Small non-coding RNAs (sRNAs) are important regulators of gene expression in bacteria, including cyanobacteria (21). Most frequently they bind to their target mRNAs close to the ribosome binding site therefore inhibiting translation, but a large diversity of alternative regulatory mechanisms by sRNAs are known (22). They are involved in the regulation of diverse processes related to response to stress or adaptation to a changing environment, included cell differentiation. In Bacillus several sRNAs have been identified that are sporulation-specific and with expression restricted to the forespore (23-26). In heterocyst-forming cyanobacteria sRNAs and antisense RNAs have been identified with heterocyst-specific expression (27-29). A heterocyst-specific antisense RNA is involved in carbon fixation shutdown in heterocysts (30). NsiR1 is a small RNA of about 60 nucleotides encoded in 12 tandem copies in the region upstream of hetF in Nostoc sp. PCC 7120 (27) (Fig. 1A). NsiR1 is expressed specifically in cells becoming heterocyst before any sign of morphological differentiation is observed (31). The different copies are transcribed from individual promoters that contain the DIF1 sequence motif associated to heterocyst-specific transcription (10, 28). Central copies nsiR1.3 to nsiR1.9 are identical whereas the other copies, located at the periphery of the array, have some sequence divergence. According to the position of the TSS of hetF (10, 27), nsiR1.1 overlaps the hetF 5’ UTR. NsiR1.1 could therefore function as an antisense of the hetF transcript. As the seven identical copies (3-9, thereafter nsiR1.4) are very similar to nsiR1.1, they could also potentially regulate the expression of hetF or other targets in Nostoc sp. PCC 7120. In this work we show that Nostoc strains constitutively overexpressing nsiR1.1 or nsiR1.4 have distinct phenotypes related to heterocyst differentiation, indicating that they have non-overlapping targets. NsiR1.1 negatively regulates hetF, and NsiR1.4 negatively regulates the hetP homolog alr3234. Therefore, sequence divergence between the different versions of NsiR1 allows regulation of diverse aspects of heterocyst differentiation through their effects on the amounts of HetF (influencing HetR accumulation) and Alr3234 (influencing commitment) specifically in heterocysts.

Results

NsiR1 interacts with the 5’ UTR of hetF.

NsiR1.1 is transcribed antisense to the 5’ UTR of hetF in Nostoc sp. PCC 7120 and this arrangement is conserved along genomes of heterocystous cyanobacteria (27) (SI Appendix, Fig. S1 and S2), therefore we hypothesized that NsiR1.1 might regulate hetF expression. To explore this possibility the 5’-ends of hetF transcripts were analyzed by retrotranscription using an

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oligonucleotide complementary to hetF 5’ UTR. 5’-ends corresponding to the TSS of hetF, position 4273166f (10, 27) in the Nostoc sp. PCC 7120 genome, were in fact observed in RNA samples isolated from both the wild type strain and the hetR mutant growing in the presence of combined nitrogen (indicated with an asterisk in Fig. 1B). However, when the cells were subjected to nitrogen deficiency, a HetR-dependent stop appeared in a position corresponding to that of the transcriptional start site of nsiR1.1 (4273242r, black triangle in Fig. 1B). Retrotranscriptase stops when it encounters an obstacle, for example, regions of strong secondary structure or a double stranded RNA such as a sRNA-mRNA duplex (32). Our results therefore suggest that the additional HetR-dependent stop might be due to binding of NsiR1, whose expression is HetR-dependent (27, 28), to hetF 5’ UTR.

If NsiR1.1 negatively regulates hetF, we would expect less HetF protein in heterocysts than in vegetative cells because nsiR1 transcription is heterocyst-specific. To test this hypothesis, we constructed Nostoc sp. PCC 7120 strains that express, under the control of a constitutive promoter (rnpB), a translational fusion of the hetF 5’ UTR plus 60 nucleotides of its coding region to gfpmut2 (plasmid pMBA93, Fig. 1A). A strain expressing gfpmut2 and its original 5’UTR from the rnpB promoter was constructed as a control (plasmid pMBA92, Fig. 1A). As expected the strain bearing the control plasmid pMBA92 showed similar green fluorescence in vegetative cells or heterocysts (Fig. 1C). However, the strain bearing pMBA93 showed detectable green fluorescence in vegetative cells but not in heterocysts (Fig. 1C), suggesting heterocyst-specific regulation at the level of the hetF 5’ UTR.

We then tested the interaction of NsiR1.1 with hetF 5’ UTR using a heterologous reporter assay in E. coli (33) in which the 5’ UTR of hetF plus 60 nucleotides of its coding region was fused to the gene sfgfp encoding superfolder GFP. Because identical copies (nsiR1.3 to nsiR1.9) are very similar to nsiR1.1 (27) (SI Appendix, Fig. S3), we also tested the interaction between the hetF 5’ UTR and the sRNA encoded in one of the identical copies (nsiR1.4). We co- expressed in E. coli the hetF::sfgfp fusion together with NsiR1.1, NsiR1.4 or a control, unrelated RNA. Cells carrying the hetF::sfgfp translational fusion showed significant fluorescence, indicating that the translational initiation region of hetF was functional in E. coli (Fig. 1D and E). The fluorescence of cells carrying the hetF::sfgfp fusion decreased about 70% or 40% of the control when the fusion was co-expressed with NsiR1.1 or NsiR1.4, respectively, suggesting that each of these versions of NsiR1 were able to interact with the 5’ UTR of hetF (Fig. 1D and E) when transcribed in a heterologous system.

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Figure 1. NsiR1 interacts with hetF 5’ UTR. (A) Schematic representation of the genomic region encoding NsiR1 in Nostoc sp. PCC 7120 and constructs in plasmids pMBA92 and pMBA93. Bent arrows indicate the transcriptional start sites. Identical repeats of nsiR1 (3-9, nsiR1.4) are shown as red arrows. Repeats of nsiR1 with some sequence divergence (1, 2, 10, 11 and 12) are shown in light blue except repeat 1 (nsiR1.1), antisense to hetF, that is shown in green. (B) Primer extension analysis of the hetF transcript using RNA isolated from Nostoc sp. PCC 7120 wild type strain (WT) or a hetR mutant strain at different times after removal of combined nitrogen. The position of the TSS of hetF (4273166f) is indicated with an asterisk. The position corresponding to the TSS of nsiR1.1 (4273242r) is indicated with a black triangle. On the right, a sequencing reaction performed with the same oligonucleotide used for primer extension. (C) Top, confocal fluorescence images (red channel, green channel and bright field image) of filaments from strains bearing plasmids pMBA92 (control, gfp gene expressed from the rnpB promoter) or pMBA93 (translational fusion of the hetF 5’ UTR to the gfp gene expressed from the rnpB promoter) growing on top of nitrogen- free medium. Bottom, quantification of the signals for the green (GFP) and red (autofluorescence) channels. Heterocysts are indicated with numbers. (D) GFP fluorescence of E. coli cultures bearing different combinations of a plasmid expressing 5’ UTR hetF::sfgfp with plasmids expressing NsiR1.1, NsiR1.4 or an unrelated control RNA. The data are represented as mean and standard deviation of results from cultures corresponding to four independent colonies after normalization for cell density and subtraction of background fluorescence in cells bearing pXG-0. (E) FACS-based reporter assays. The plot shows the distribution of fluorescence in 10,000 E. coli cells bearing plasmids expressing 5’ UTR hetF:sfgfp and the unrelated control RNA (black), NsiR1.1 (green) or NsiR1.4 (red). The inset shows the average fluorescence of each sample. T-test P-value <0.05*;< 0.01*.

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Taken together, the results obtained in Nostoc sp. PCC 7120 and E. coli suggested a post transcriptional negative regulation exerted by NsiR1 on hetF expression. The overlapping of NsiR1.1 and the hetF 5’ UTR extends between positions 20 and 78 of the UTR, 180 nucleotides upstream the start codon. To analyze the regulatory mechanism, we carried out in vitro footprinting assays of NsiR1.1 and hetF 5’ UTR interaction (Fig. 2). 32P-labeled hetF 5’ UTR (a fragment extending from the TSS at position −257 to +60 with respect to the start of the coding sequence) was incubated with unlabeled NsiR1.1 and probed with RNase T1, RNase A, or lead(II) acetate (Fig. 2A).

Figure 2. In vitro footprinting assay of the interaction between NsiR1.1 and the hetF 5’ UTR. (A) 5’-labeled hetF 5’ UTR RNA was treated with RNase T1, RNase A, or lead(II) acetate (Pb+2) in the absence (-) or presence (+) of NsiR1.1. Samples were loaded on two gels that were run to different lengths. Vertical bars indicate the NsiR1.1 binding site (red), a region with increased sensitivity to the probes in the presence of NsR1.1 (green) and the region including the

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start codon (orange). Nucleotides with differential sensitivity to RNase T1 or RNase A in the presence or absence of NsiR1.1 are indicated with colored triangles. Nucleotide positions of hetF 5’ UTR are shown on the left. C, untreated control; OH, alkaline ladder; T1s, RNase T1 sequencing ladder. (B, C) Models of the secondary structure of hetF 5’ UTR from Nostoc sp. PCC 7120 without (B) or with bound NsiR1.1 (C). Nucleotides corresponding to the areas highlighted by vertical bars in the gels are colored in the same way. Nucleotides with differential sensitivity to RNase T1 or RNase A outside the NsiR1 binding site are indicated with colored triangles. Start codon is in purple with a purple frame.

A footprint was detected between positions 23 and 81 of hetF 5’ UTR (Fig. 2A, red bar), in agreement with the perfect antisense sequence complementary to NsiR1.1. In addition to this footprinting, we also detected some structural changes when hetF 5’ UTR was combined with NsiR1.1; a region between positions 98 and 104 of hetF 5’ UTR became more sensitive to RNase T1, RNase A and lead(II) acetate (Fig. 2A, green bar) and the region including the start codon became more protected (Fig. 2A, orange bar). These results indicate that upon binding of NsiR1 there is a structural reorganization of the 5’ UTR of hetF resulting in reduced access to the initiation codon (Fig. 2B and C) that could explain the reduction of HetF expression in heterocysts.

Strains with altered levels of different versions of NsiR1.

We have constructed strains with constitutive high levels of NsiR1.1 (OE_NsiR1.1) or NsiR1.4 (OE_NsiR1.4) transcribed from the strong trc promoter to the T1 terminator of the rrnB gene of E. coli (Fig. 3A). In addition, because we have introduced an NsiI restriction site (6 bp) at the 5’ end of nsiR1.1 or nsiR1.4 to facilitate cloning, this 6 nt tag allows the discrimination, based on its length, of molecules of NsiR1.1 and NsiR1.4 transcribed from their native promoters from molecules expressed from the trc promoter in the constructs introduced in Nostoc sp. PCC 7120.

The location of nsiR1.1 in the 5’ UTR of hetF precludes the deletion of all copies of nsiR1 without removing a segment of hetF 5’ UTR. To deplete NsiR1 without altering the genomic region upstream of hetF we have followed a strategy similar to (34) and generated a Nostoc strain (OE_as_NsiR1) that constitutively expresses the sequence complementary to nsiR1.4 from the trc promoter (Fig. 3A). This antisense RNA would act as a sponge neutralizing the accumulation of nsiR1.

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Figure 3. Nostoc sp. PCC 7120 strains with altered levels of NsiR1. (A) Scheme of DNA fragments cloned in the plasmids used to generate the different strains. Transcription start site (bent arrows), Rho-independent terminator of NsiR1 (small stem loop), T1 terminator (large stem loop), trc promoter, probe for nsiR1.1 (green line), probe for nsiR1.4 (red line), oligonucleotide used for strand-specific probe labeling (black triangle) and sequences corresponding to nsiR1.1 (green arrow) and nsiR1.4 (red arrows) are indicated. (B) Northern blots using RNA extracted from the different strains at different times after nitrogen removal hybridized with probes for NsiR1.1 (top), NsiR1.4 (middle) and 5S rRNA (bottom) as a loading control. Endogenous NsiR1 (black triangles) and the three RNA species constitutively expressed from the trc promoter, NsiR1.1 (green triangle), NsiR1.4 (red triangle) and as_NsiR1 (grey triangles), are indicated. (C) Primer extension analysis of hetF transcript with RNA isolated from the mutant strains at different time points after combined nitrogen removal. The position of the TSS of hetF (4273166f) is indicated with an asterisk. The position corresponding to the TSS of nsiR1.1 (4273242r) is indicated with a black triangle. Stops provoked by NsiR1.1 (green triangle) and NsiR1.4 (red triangles) constitutively expressed from the trc promoter are indicated. On the right, a sequencing reaction performed with the same oligonucleotide used for primer extension. (D) Accumulation of HetR in the mutant strains 18 h after combined nitrogen removal. Western blots were carried out with proteins extracted from three independent cultures of each strain using antibodies against HetR or GroEL (used as loading control). (E) Quantification of Western blot shown in (D). The data are presented as the mean ± standard deviation of the HetR signal normalized to the GroEL signal of the three independent clones of each strain shown in (D). The mean of HetR accumulation of the OE_C strain in each gel is used as 100%. **T-test P-value < 0.01.

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Using probes specific for NsiR1.1 or NsiR1.4 we have verified by Northern blot the constitutive expression of NsiR1.1 and NsiR1.4 under the control of the trc promoter in strains OE_NsiR1.1 and OE_NsiR1.4, respectively (Fig. 3B). In addition, Northern blot hybridization also showed that, in the OE_C strain, signals corresponding to transcription of endogenous nsiR1.4 were stronger than those corresponding to nsiR1.1, as expected from the different number of copies of nsiR1.4 (seven copies) versus nsiR1.1 (single copy). Transcripts corresponding to NsiR1.1 and NsiR1.4 whose transcription did not stop at their endogenous Rho-independent terminators but instead were terminated at the T1 terminator were also observed (about 100 nt long). Regarding the OE_as_NsiR1 strain, Northern blot hybridization showed that the accumulation of NsiR1.1 and NsiR1.4 were diminished in this strain, suggesting effective removal of endogenous native NsiR1 by overexpression of the antisense sponge (Fig. 3B). Both probes used included the T1 terminator, present in as_NsiR1 (Fig. 3A), what allowed its detection in the OE_as_NsiR1 strain (Fig. 3B).

We were unable to quantify hetF expression by Northern blot or RT-PCR, suggesting that, similar to the observations previously made in Nostoc punctiforme (17), the hetF transcript is likely unstable or accumulates in small quantity in Nostoc sp. PCC 7120. We have instead analyzed the accumulation of hetF transcripts and their putative interaction with the different versions of NsiR1 by primer extension assays (Fig. 3C). Similar to the observation made in the case of the wild-type strain (Fig. 1B), extension products that stopped at the position corresponding to the TSS of the chromosomal nsiR1.1 were observed in the OE_C, OE_NsiR1.1 and OE_NsiR1.4 strains when the cells were subjected to nitrogen deficiency (Fig. 3C, black triangles). In the OE_as_NsiR1 strain, overexpressing a sponge that is expected to retire endogenous NsiR1, these stops are only observed after 12 hours of nitrogen deficiency an in reduced amounts, consistent with the observation that the sponge RNA is efficiently removing NsiR1 in this strain (Fig. 3B). As expected, and in contrast to the other three strains included in the experiment, strain OE_NsiR1.1 showed no full-length extension products that corresponded to the 5’-end of hetF. Instead, a strong stop around the start position of nsiR1.1 was observed irrespective of nitrogen removal (Fig. 3C, green triangle), suggesting that the presence of NsiR1.1 expressed from the constitutive trc promoter was responsible for such stops. Similarly, stops independent of the nitrogen condition but weaker than those observed in OE_NsiR1.1 were also observed in the OE_NsiR1.4 strain (Fig. 3C, red triangles). Taken together, these observations indicate that the HetR-dependent NsiR1.1 sRNA (and possibly NsiR1.4) interacts with the 5’ UTR of hetF in Nostoc sp. PCC 7120.

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Based on data from fluorescence microscopy in Nostoc strains bearing pMBA93, the observations made in E. coli, and in vitro footprinting assays (Fig. 1 and 2), NsiR1.1 and, to a lesser extent, NsiR1.4 could exert a negative regulation of hetF expression. It is known that a hetF mutant accumulates more HetR protein than the WT strain (18). We therefore quantified the accumulation of HetR in the OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains 18 h after combined nitrogen removal (Fig. 3D and E). Using three independent clones of each strain, we determined that strain OE_NsiR1.1 accumulated about twice more HetR protein than the OE_C strain. OE_NsiR1.4 also seems to accumulate more HetR protein than the OE_C strain, but in this case, perhaps due to the effects of NsiR1.4 on HetF accumulation being weaker, the results were not statistically significant. In order to analyze whether the increased amounts of HetR protein could be instead due to increased transcription of hetR in the overexpressing strands, we measured the accumulation of hetR transcripts in all four strains analyzed above (OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1) (SI Appendix, Fig. S4). After 18 h of nitrogen deficiency the amounts of hetR transcript in the OE_NsiR1.1 strain were not above the amounts observed in the control strain (OE_C), indicating that the increased amount of HetR in strain OE_NsiR1.1 was not the consequence of increased transcription of the hetR gene.

Phenotype of OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains.

In order to analyze the effects of altered NsiR1 expression at the phenotypic level we tested growth of the OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains in solid media containing different nitrogen sources (Fig. 4A). All four strains grew similarly in the presence of ammonium, however strain OE_NsiR1.4 grew worse in the absence of combined nitrogen and a bit slower in the presence of nitrate than the rest of strains. In liquid cultures, strain OE_NsiR1.4 showed slight phenotypic differences in the presence of ammonium. Some cells were larger and with some division problems. The differences were more dramatic after combined nitrogen removal (SI Appendix, Fig. S5). After nitrogen removal, strain OE_NsiR1.4 differentiated cells that were stained by Alcian blue (indicative of the presence of heterocyst polysaccharides) but some filaments had enlarged cells which resembled the phenotype previously described for the hetF mutant (18). OE_NsiR1.1, also contained some filaments with larger cells under nitrogen deprivation (SI Appendix, Fig. S5).

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Figure 4. Phenotype of OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains. (A) Growth of solid media. Cells from liquid cultures of the indicated strains grown in the presence of nitrate were collected and resuspended in BG110 at an OD750= 0.17. Five-fold serial dilutions were prepared and 10 µl of each dilution plated on BG110 plates containing + - ammonium (NH4 ), nitrate (NO3 ) or lacking nitrogen (N2). Pictures were taken after 10 days of incubation at 30ºC. (B) Bright field images of OE_C, OE_NsiR1.1 and OE_NsiR1.4 strains growing on BG110 plates containing nitrate. Heterocysts are indicated with black triangles. (C) Nitrate and nitrite reductase assays. The data are given as the mean ± standard deviation of nitrate or nitrite reductase specific activity (mU/mg total protein) of three independent clones of the OE_C strain and two independent clones of OE_NsiR1.4 in different nitrogen sources; ND, not determined. (D) Northern blots with RNA isolated from the indicated strains at different time points (indicated in hours) after nitrogen removal and hybridized with probes for nifH (upper panels) and rnpB (bottom panels) as a loading control. Sizes are indicated on the left in kb.

The slightly slower growth of strain OE_NsiR1.4 in solid media in the presence of nitrate (Fig. 4A) prompted us to analyze the filaments in more detail, leading to the observation that OE_NsiR1.4 contained patterned heterocysts in the presence of nitrate in solid media (Fig. 4B). To verify that the presence of heterocysts in strain OE_NsiR1.4 was not due to its inability to assimilate nitrate, we measured nitrate and nitrite reductase activities (Fig. 4C). Although strain OE_NsiR1.4 showed about half of the nitrate and nitrite reductase activities of OE_C

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strain, a previously described mutant strain with a similar reduction in nitrite reductase activity does not differentiate heterocysts (35, 36), therefore suggesting that the heterocyst differentiation we observe in strain OE_NsiR1.4 in the presence of nitrate is not due to a defect in nitrate assimilation.

We also analyzed the timing of expression of nifH during heterocyst differentiation upon nitrogen removal in the different strains (Fig. 4D). OE_NsiR1.1 and OE_NsiR1.4 showed no difference with respect to the control, however OE_as_NsiR1 showed a delay in the accumulation of nifH mRNA.

NsiR1.4 interacts with alr3234 5’ UTR.

Overexpression of NsiR1.1 (OE_NsiR1.1) or NsiR1.4 (OE_NsiR1.4) leads to different phenotypes (Fig. 4), suggesting that, although they both can regulate hetF to a different degree, they might also regulate additional non-overlapping targets.

A previous computational prediction (27) identified alr3234 as a possible target of NsiR1.4. The predicted interaction of NsiR1.4 was located in the translation initiation region of the alr3234 mRNA (Fig. 5A). IntaRNA (37) also predicted an interaction between NsiR1.1 and the 5’ UTR of alr3234 but the hybridization energy was much lower (Fig. 5A). We performed in vitro footprinting assays of the interaction of NsiR1.1 and NsiR1.4 (Fig. 5B) with the wild-type version of alr3234 5’ UTR, or with a version bearing two point mutations, alr3234 5’ UTR (MUT), that should interact more efficiently with NsiR1.1 than with NsiR1.4 (Fig. 5A). The 32P-labeled alr3234 5’ UTR versions were incubated with unlabeled NsiR1.1 or NsiR1.4 and probed with RNase T1 or lead(II) acetate (Fig. 5B). We only detected footprints in the predicted region when alr3234 5’ UTR (WT) was combined with NsiR1.4 or when alr3234 5’ UTR (MUT) was combined with NsiR1.1 (Fig. 5B).

We then tested the interaction of NsiR1.4 and NsiR1.1 with the two versions of the alr3234 5’ UTR in the heterologous reporter system in E. coli. Cells carrying the alr3234::sfgfp fusion did not show significant fluorescence (not shown) but we could instead measure the accumulation of Alr3234::sfGFP in E. coli by Western blot with anti-GFP antibodies (Fig. 5C). The accumulation of Alr3234::sfGFP translated from alr3234 5’ UTR (WT) decreased about 70% with respect to the control strain when NsiR1.4 was co-expressed and about 30% when NsiR1.1 was co-expressed (Fig. 5C and D). However, the accumulation of Alr3234::sfGFP translated from alr3234 5’ UTR (MUT) equally decreased when this mRNA was co-expressed with NsiR1.1 or NsiR1.4 (Fig. 5C and D).

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Figure 5. NsiR1 interacts with alr3234 5’ UTR. (A) Predicted interaction according to IntaRNA between NsiR1.1 or NsiR1.4 and two versions of the 5’ UTR of alr3234 (WT and MUT). Nucleotides corresponding to NsiR1.1, NsiR1.4, start codon of alr3234 and mutations introduced in the alr3234 5’ UTR MUT version are shown in green, red, blue and orange, respectively. (B) RNase T1 and lead(II) acetate (Pb+2) footprinting of the interaction between NsiR1.1 or NsiR1.4 with alr3234 5’ UTR. The protected areas are indicated by red vertical bars. Nucleotide positions of alr3234 5’ UTR are shown on the left. C, untreated control; OH, alkaline ladder; T1s, RNase T1 sequencing ladder. (C) Accumulation of GFP protein in E. coli cells bearing several combinations of plasmids expressing different versions of alr3234::sfgfp and control RNA (C), NsiR1.1 (#1) or NsiR1.4 (#4). Western blots were carried out using antibodies against GFP or GroEL (loading control). (D) Quantitation of Western blots. The data are presented as the mean ± standard deviation of GFP signal normalized to GroEL signal of three independent experiments like the one shown in (C) with each plasmid combination. The mean of GFP accumulation of the control strain in each experiment is used as 100%. T-test P-value <0.05*;< 0.01*; 0.001***.

We constructed a Nostoc strain bearing, under the control of the constitutive rnpB promoter, a translational fusion of the alr3234 5’ UTR plus 60 nucleotides of its coding region to gfpmut2 (pMBA94). The strain bearing pMBA94 showed low but detectable green fluorescence

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in vegetative cells but not in mature heterocysts (Fig. 6A), in agreement with heterocyst-specific post transcriptional regulation by NsiR1. The reduction in green fluorescence of the alr3234::sfgfp fusion in heterocysts occurred even in immature heterocysts at an early stage of differentiation that still had red autofluorescence (Fig. 6A).

The expression of alr3234 is very low irrespective of the nitrogen source (20) precluding accurate quantitation of mRNA changes. However, it has been described that in an alr3234 mutant pro-heterocysts commit to differentiation earlier than in the wild type strain (20). Therefore, to determine possible changes in the commitment time among the strains with altered levels of NsiR1, we carried out a heterocyst-commitment assay in which ammonium is added at different time points after nitrogen removal and the final frequency of heterocysts is recorded. Whereas all four strains analyzed showed a similar frequency of heterocyst after 48 hours of nitrogen removal (5-7%), the heterocyst-commitment assay showed that strain OE_NsiR1.4 had an earlier onset of commitment than the OE_C strain (Fig. 6B), in agreement with heterocyst-specific post transcriptional regulation of alr3234 by NsiR1.4. A slight delay in commitment was observed in strain OE_as_NsiR1 (Fig. 6B). In addition, and opposite to OE_NsiR1.4, strain OE_NsiR1.1 had a significant delay in commitment.

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Figure 6. Post-transcriptional regulation of alr3234 in Nostoc sp. PCC 7120 and commitment assay in strains with altered levels of NsiR1. (A) Top, confocal fluorescence images (red channel, green channel and bright field image) of filaments growing on top of nitrogen-free medium and carrying under the control of the rnpB promoter the gfp gene (plasmid pMBA92) (left) or a translational fusion of the alr3234 5’ UTR plus 60 nucleotides of the coding sequence to the gfp gene (plasmid pMB94) (right). Bottom, quantification of the signals for the green (GFP) and red (autofluorescence) channels. Heterocysts are indicated with numbers. (B) Determination of time of heterocyst commitment. After combined nitrogen removal (t=0h), ammonium was added at the indicated times and the frequency of heterocyst was determined 24-36 h after ammonium addition. For each strain the data are presented as the mean and standard deviation of two cultures from two independent clones.

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Discussion

Heterocyst differentiation is a complex process that requires precise regulatory mechanisms in order to establish exclusive transcriptional programs for different cell types (vegetative cells, immature pro-heterocysts and mature heterocysts). Heterocyst differentiation is triggered by mutual induction of expression of the transcriptional regulator NtcA and the master regulator HetR in vegetative cells becoming heterocysts (7, 8, 38). The accumulation of active HetR in pro-heterocysts seems to trigger the downstream cascade. NsiR1 was the first HetR-dependent sRNA identified in Nostoc sp. PCC 7120 (27), but it is also well conserved in non- heterocyst-forming filamentous cyanobacteria (29). However, while most non-heterocyst strains contain a single copy of NsiR1, heterocystous strains have multiple tandem copies with slight sequence divergence (SI Appendix, Fig. S1, Fig. S2). The increase in the number of copies of NsiR1, that correlates with the acquisition of heterocysts in evolution, and their sequence divergence, could allow some degree of functional specialization by regulating a variety of targets.

According to the position of the TSS of hetF in Nostoc sp. PCC 7120 (10, 27), the nsiR1 copy closest to hetF (nsiR1.1) could function as an antisense of hetF 5’ UTR. This function of NsiR1.1 may be conserved in heterocyst-forming cyanobacteria because most of them have a nsiR1 homolog in the upstream region of hetF in antisense orientation (SI Appendix, Fig. S2).

Co-expression experiments in E. coli indicate that NsiR1.1, and to a lower extent NsiR1.4, can repress the expression of hetF (Fig. 1D and E). A translational fusion of hetF 5’ UTR to GFP had reduced expression in heterocysts (Fig. 1C), as expected if its translation is repressed by NsiR1, that is expressed at an early stage during heterocyst development (31). In addition, primer extension analysis in Nostoc sp. PCC 7120 also suggested a putative interaction of NsiR1.1 with hetF 5’ UTR based on the stop of the retrotranscriptase (Fig. 1B). Two observations point to post-transcriptional regulation of translation rather than transcriptional interference as the mechanism for hetF regulation by NsiR1. First, negative regulation of hetF expression is observed in the E. coli system, in which NsiR1 and the hetF 5’ UTR are transcribed from E. coli promoters in separate plasmids. Second, the observation that NsiR1.1, which perfectly matches the hetF 5’ UTR, and NsiR1.4 which contains several nucleotide changes, repress hetF translation to a different extent (Fig. 1D), is indicative of post-transcriptional regulation by perfect (NsiR1.1) or imperfect (NsiR1.4) base pairing. Finally, the quantity of hetF mRNA did not significantly decrease when NsiR1 was present (Fig. 1B), indicating that NsiR1 did not affect hetF transcription.

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The interaction site of NsiR1 with the hetF 5’ UTR was located relatively far upstream from the hetF start codon, precluding repression of translation by direct interaction of NsiR1 with the translation initiation region and pointing to a translational regulation based on a conformational change of the hetF 5’ UTR. In vitro footprinting data and computational analysis showed that the hetF 5’ UTR is highly structured and the interaction with NsiR1.1 provokes a conformational change in which the region of the start codon of hetF gets more protected (Fig. 2), providing a mechanistic explanation to the negative regulation of hetF expression by NsiR1.1.

In order to study the function of NsiR1, we created strains of Nostoc sp. PCC 7120 with altered levels of different versions of NsiR1 (Fig. 3). The overexpression of NsiR1.1 or NsiR1.4 under the control of the trc promoter was irrespective of the nitrogen source and strong enough to overpass the endogenous expression of NsiR1, that was induced upon nitrogen removal. The OE_as_NsiR1 strain had reduced amounts of NsiR1, implying that the interaction between NsiR1 and its antisense results in its degradation (Fig. 3B). Primer extension assays with RNA extracted from strains with altered levels of different versions of NsiR1 confirmed the interaction of NsiR1.1 and NsiR1.4 with hetF 5’ UTR (Fig. 3C). Expression of hetF is low and the transcripts are unstable (17, 18), therefore we were unable to quantify changes in hetF expression in our mutant strains. Instead, because one of the features described for a hetF mutant is the accumulation of higher levels of HetR protein (18), we analyzed HetR protein levels as an indication of HetF function. In fact, strains OE_NsiR1.1 and, to a lesser extent, OE_NsiR1.4, had higher amounts of HetR, as expected for a strain that would have less HetF (Fig. 3D and E). This higher amount of HetR was not due to increased hetR transcription in strains OE_NsiR1.1 and OE_NsiR1.4 (SI Appendix, Fig. S4). Finally, OE_NsiR1.1 had a phenotype partially similar in some aspects to that described for a hetF mutant. A hetF mutant grows poorly in media without combined nitrogen, has enlarged cells and cannot differentiate heterocysts (18). OE_NsiR1.1 showed some filaments with larger cells in liquid cultures growing without combined nitrogen (SI Appendix, Fig. S5) and was delayed in the timing of commitment (Fig. 6B). Taken together, all these in vivo and in vitro results strongly support a role for NsiR1 in reducing the amount of HetF in cells becoming heterocysts.

A computational analysis (27) identified alr3234, a hetP homolog described as a negative modulator of heterocyst commitment (20), as a potential target of NsiR1.4. We confirmed the interaction between NsiR1.4 and alr3234 mRNA by in vitro footprinting and coexpression in E.coli (Fig. 5). NsiR1.4 binds to a region that includes the start codon of alr3234, and thus could inhibit translation initiation by hampering ribosome access. One of the phenotypic features described for an alr3234 mutant is earlier commitment to heterocyst

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differentiation (20). Timing of commitment in OE_NsiR1.4 was in fact shifted to earlier time points (about 2 hours earlier than OE_C) while OE_as_NsiR1 had a slight delay in commitment (Fig. 6B), supporting an inhibitory effect of NsiR1.4 on alr3234. The putative positive effect of NsiR1.4 on heterocyst commitment is also in line with the observation that OE_NsiR1.4 differentiates heterocyst even in the presence of nitrate (Fig. 4B). Alr3234 represents a brake that delays commitment, providing a lag time to avoid unnecessary differentiation under fluctuating conditions (20). We hypothesize that only when enough NsiR1 has accumulated as a consequence of constant nitrogen deficiency is the amount of Alr3234 reduced and the brake on commitment released.

Despite the slight difference in sequence between NsiR1.1 and NsiR1.4 (SI Appendix, Fig. S3), strains OE_NsiR1.1 and OE_NsiR1.4 had different phenotypes. They grew differentially in solid media with different nitrogen sources (Fig. 4A). OE_NsiR1.4, but not OE_NsiR1.1, differentiated heterocysts with a normal pattern in solid media with nitrate as nitrogen source, a condition that suppresses heterocyst development in the wild type (Fig. 4B). Finally, these strains differ in the timing of commitment to heterocyst differentiation (Fig. 6B). Thus, these results suggest a divergence in the functions of NsiR1.1 and NsiR1.4, that could be explained by their different effects on hetF and alr3234 or by interactions with additional unknown targets specific for each of them. The positive effect of NsiR1.4 on differentiation was in accordance with the observation that OE_as_NsiR1 (a strain with a lower amount of the NsiR1 pool, mostly formed by NsiR1.4) was delayed in differentiation (Fig. 4D) and slightly delayed in commitment (Fig. 6B).

In this work we describe how different versions of one sRNA regulate different targets. Other instances of sRNAs encoded by multiple repeats have been described in diverse bacteria (39). In most cases there is a regulatory redundancy and the different copies of one given sRNA act additively to regulate one particular target. In other cases they are non-redundant, regulating different targets. NsiR1.1, on one side, and the identical copies NsiR1.3-NsiR1.9 on the other, represent divergent versions of a sRNA with partially redundant targets but possibly also different targets.

Present models of events leading to heterocyst differentiation and patterning are based on transcriptional regulation and protein-protein interactions. Beyond those mechanisms, we here identify a role for a sRNA that specifically accumulates in heterocysts. We propose that, even though hetF and alr3234 are transcribed constitutively, upon nitrogen deprivation the early and specific expression of NsiR1.1 and NsiR1.4 in pro-heterocysts would

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provoke the post-transcriptional repression of hetF and alr3234, resulting on a local accumulation of HetR in pro-heterocyst (triggering the downstream cascade of heterocyst differentiation) and releasing the brake of alr3234 inhibition of the commitment of pro- heterocyst into differentiation (Fig. 7). So, the specific post-transcriptional regulation exerted by different versions of NsiR1 in the multicellular cyanobacterium Nostoc sp. PCC 7120 is a good example of how complexity can emerge from the superposition of simple patterns, constitutive transcription (hetF, alr3234) plus a cell-type specific negative post-transcriptional regulation mediated by a small RNA (NsiR1). Further study of additional potential targets for NsiR1, and the characterization of additional heterocyst-specific non-coding RNAs (28) could reveal the importance of these molecules in the regulation of this developmental process.

Figure 7. Model of the function of NsiR1.1 and NsiR1.4 in the regulation of heterocyst differentiation and commitment. Transcriptional regulation is indicated by black arrows. Negative post transcriptional regulation by NsiR1 is indicated by red lines with blunt ends. HetF regulation of HetR is indicated by blue lines. Thin lines indicate weaker regulation than thick lines. Different letter size portraits the different relative amounts of the proteins or RNAs displayed in the two types of cells. Question marks indicate possible additional targets of NsiR1.1 and NsiR1.4. Green dashed line delimits events occurring in heterocysts (left) from events occurring in vegetative cells (right).

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Materials and Methods

Strains and growth conditions.

The different Nostoc sp. PCC 7120 strains used in this work (SI Appendix,Table S1) were grown in BG11 medium (40) as detailed in SI Appendix, SI Materials and Methods. Escherichia coli strains (SI Appendix, Table S1) were grown in LB medium, supplemented with appropriate antibiotics (41).

Construction of Nostoc sp. PCC 7120 derived strains.

Details on the construction of plasmids (SI Appendix, Tables S2) are given in SI Appendix, SI Materials and Methods.

In vitro synthesis and labelling of RNA.

RNAs were in vitro transcribed (MEGAscript Kit; Life Technologies, #AM1330) from PCR-generated DNA templates using the primers indicated in SI Appendix, Table S3. Details of the transcription, purification and labeling procedures are given in SI Appendix, SI Materials and Methods.

In vitro structure probing and footprinting.

RNase T1, RNase A and lead(II) acetate were used for structure probing with in vitro transcribed RNAs (SI Appendix, Table S4) essentially as described (42). Details of the procedures are given in SI Appendix, SI Materials and Methods

RNA isolation, Northern blot analysis and primer extension assays.

Total RNA was isolated using hot phenol as described (43) with modifications (29). Strand specific 32P probe labeling and Northern blots were performed as detailed in SI Appendix. Primer extension of 5´ ends of hetF was performed as previously described (44) using oligonucleotide 105 (SI Appendix, Table S3) labeled with [ -32P] ATP and 10 µg of total RNA.

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Heterocyst differentiation and commitment assay.

Heterocysts were counted after staining with Alcian blue as described (45). Number of heterocysts and commitment time were determined as described in SI Appendix, SI Materials and Methods.

Nitrate and nitrite reductase assay.

Nitrate and nitrite reductase were performed as described (46, 47). More details in SI Appendix, SI Materials and Methods.

Western blot analysis.

For HetR quantitation in Nostoc sp. PCC 7120 filaments, protein extracts were prepared using glass beads and Western blots were performed as detailed in SI Appendix, SI Materials and Methods. Antibodies against Nostoc sp. PCC 7120 HetR (48) and E.coli GroEL (Sigma-Aldrich) were used. Quantitation of alr3234::GFP in E. coli cells was performed as detailed in SI Appendix, SI Materials and Methods. Antibodies against GFP (monoclonal, Roche) and E. coli GroEL (Sigma-Aldrich) were used.

Reporter assay for in vivo verification of targets.

We used the GFP reporter system described in (49) with the superfolder GFP (sfGFP) and plasmid pXG10-SF (33) for the experimental target verification in E.coli. The construction of the different GFP fusions used (SI Appendix, Table S5) and site directed mutagenesis are described in SI Appendix, SI Materials and Methods.

Fluorescence microscopy.

Fluorescence of Nostoc sp. PCC 7120 filaments carrying plasmid pMBA92, pMBA93 and pMBA94 (SI Appendix, Table S6) growing on top of solidified nitrogen-free medium, was analyzed and quantified as described (31) using a Leica HCX PLAN-APO 63X 1.4 NA oil immersion objective attached to a Leica TCS SP2 confocal laser-scanning microscope. Samples were excited at 488 nm by an argon ion laser and the fluorescent emission was monitored by collection across windows of 500–538 nm (GFP) and 630–700 nm (cyanobacterial autofluorescence).

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Computational and statistical methods.

Computational and statistical methods are described in detail in SI Appendix, SI Materials and Methods.

Acknowledgements

This work was supported by grants BFU2013-48282-C2-1 from Ministerio de Economía y Competitividad, and BFU2016-74943-C2-1-P from Agencia Estatal de Investigación (AEI), Ministerio de Economía, Industria y Competitividad, both cofinanced by the European Regional Development Fund, to A.M.P. M.B.A is the recipient of a predoctoral contract from Ministerio de Educación, Cultura y Deporte, Spain (FPU014/05123 and EST16-00088). We are grateful to Wolfgang R. Hess (Freiburg University, Germany) for valuable comments on the manuscript.

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Supplementary Information for

NsiR1, a small RNA with multiple copies, regulates heterocyst differentiation and commitment in Nostoc sp. PCC 7120

Authors: Manuel Brenes-Álvarez, Marina Mingueta, Agustín Vioque1, and Alicia M. Muro-Pastor

Authors affiliation: Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones Científicas and Universidad de Sevilla, Sevilla, Spain.

aPresent address: Centro de Biología Molecular Severo Ochoa, CSIC-UAM, Nicolás Cabrera 1, 28049, Madrid, Spain.

1Corresponding Author: Agustín Vioque, Instituto de Bioquímica Vegetal y Fotosíntesis, Américo Vespucio 49, 41092, Sevilla, Spain

Phone: +34-954489519 Email: [email protected]

This PDF file includes:

Materials and Methods Figures S1 to S5 Tables S1 to S6 SI References

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Supplementary Materials and Methods

Strains and growth conditions

For Northern blot and primer extension analysis, cultures of wild-type, hetR mutant 216 (1) and different derivative strains of Nostoc sp. PCC 7120 (SI Appendix,Table S1) were

bubbled with an air/CO2 mixture (1% v/v) and grown photoautotrophically at 30ºC in BG11

medium (2) lacking NaNO3 but containing 6 mM NH4Cl, 10 mM NaHCO3, and 12 mM N-tris

(hydroxymethyl) methyl-2-aminoethanesulfonic acid-NaOH buffer (pH 7.5) (BG110C + 6 mM

NH4Cl). Nitrogen deficiency was induced by filtering, washing and resuspending cells in nitrogen-

free-medium with 10 mM NaHCO3 (BG110C). Chlorophyll concentration of the cultures was measured as described (3).

For Western blot analysis, cultures of different derivative strains of Nostoc sp. PCC 7120 (SI Appendix, Table S1) were grown in flasks at 30ºC in liquid BG11 medium for one week. Nitrogen deficiency was induced as described above.

OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains were grown in the presence of streptomycin (Sm) and spectinomycin (Sp), 2 µg/mL each (liquid medium) or 5 µg/mL each (solid medium).

Escherichia coli strains (SI Appendix, Table S1) were grown in LB medium, supplemented with appropriate antibiotics (4).

Construction of Nostoc strains with altered levels of NsiR1

Plasmids and oligonucleotides used in this work are described in SI Appendix, Table S2 and SI Appendix, Table S3.

We have used pMBA37 (5) as a backbone for overexpressing different copies of nsiR1 or an antisense to nsiR1. pMBA37 contains the trc promoter and the T1 terminator from E. coli rrnB gene. The trc promoter from this plasmid was amplified using oligonucleotides 573 and 574 (fragment 1). This product and oligonucleotide 573 were later used for PCRs with oligonucleotides 757, 758 or 512 generating fragments in which the trc promoter was fused respectively to nsiR1.1 (fragment 2), nsiR1.4 (fragment 3) and as_nsiR1 (fragment 4). After digestion of fragments 2-4 with ClaI and XhoI in the sites provided by oligonucleotides 573, 757, 758 and 512, these fragments were cloned into ClaI-XhoI digested pMBA37 vector, rendering respectively pMBA77, pMBA78 and pMBA42 (SI Appendix, Table S2).

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pMBA77, pMBA78 and pMBA42 were introduced in Nostoc sp. PCC 7120 wild type by conjugation (6) generating strains OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1, respectively (SI Appendix, Table S1).

Construction of Nostoc strains with 5’UTR of hetF and alr3234 fused to gfp

We have used pMBA91, a derivative of pMBA20 (5), as a backbone for the constitutive overexpression from rnpB promoter of a translational fusion to GFP of the 5’UTRs of hetF, alr3234 or its own 5’UTR (used as control). This vector is similar to pMBA20 but with an Nm resistance cassette instead of a SmSp resistance cassette. The Nm resistance cassette from pRL278 (7) was amplified using oligonucleotides 892 + 893. After digestion with BamHI, this product was cloned in BamHI pMBA20 vector, rendering pMBA91 (SI Appendix, Table S2).

The hetF 5’UTR was amplified using as template genomic DNA and oligonucleotides 555 + 895. gfpmut2 gene was amplified using as template pSAM270 (8) and oligonucleotides 896 + 894. A PCR product with a translational fusion between the hetF 5’UTR plus sequences encoding the first 20 amino acids of HetF and gfpmut2 was amplified using as templates the two above-described PCR fragments and oligonucleotides 555 + 894. The PCR product was digested with NsiI and XhoI and cloned in NsiI-XhoI digested pMBA91, rendering pMBA93.

The alr3234 5’UTR was amplified using as template genomic DNA and oligonucleotides 553 + 897. gfpmut2 gene was amplified using as template pSAM270 (8) and oligonucleotides 898 + 894. A PCR product with a translational fusion between the alr3234 5’UTR plus sequences encoding the first 20 amino acids of Alr3234 and gfpmut2 was amplified using as templates the two above-described PCR fragments and oligonucleotides 553 + 894. The PCR product was digested with NsiI and XhoI and cloned in NsiI-XhoI digested pMBA91, rendering pMBA94. The 5’UTR of gfpmut2 was used as control.

The gfpmut2 gene plus its 5’UTR were amplified using as template pSAM270 (8) and oligonucleotides 894 + 899. The PCR product was digested with NsiI and XhoI and cloned in NsiI- XhoI digested pMBA91, rendering pMBA92 (SI Appendix, Table S2). pMBA92, pMBA93 and pMBA94 were introduced in Nostoc sp. PCC 7120 by conjugation (6). Sequences of inserts in plasmids expressing GFP translational fusions in Nostoc sp. PCC 7120 are shown in SI Appendix, Table S6.

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In vitro synthesis and labeling of RNA

RNA transcripts were generated in vitro with MEGAscript High Yield Transcription Kit (AM1333, Ambion). The DNA templates for the transcription of hetF 5’UTR, alr3234 5’UTR (WT or MUT variant), NsiR1.1 and NsiR1.4 were generated by PCR with a primer that includes a T7 promoter sequence upstream the 5’-end of the coded RNA, and a primer matching the 3’-end of the RNA (SI Appendix, Table S3). The templates used for these PCR amplifications were genomic DNA (for hetF 5’UTR), pMBA86 (for alr3234 wild type 5’UTR), pMBA87 (for alr3234 MUT 5’UTR) or only the overlapping PCR oligonucleotides (for NsiR1.1 and NsiR1.4). The sequences of the RNAs transcribed in vitro are shown in SI Appendix, Table S4. After in vitro transcription, RNAs were treated with DNase I and purified by phenol, phenol/chloroform and chloroform extractions, ethanol-precipitated at −20 °C, and washed with 70% ethanol. 100 pmol of RNAs were labeled at the 5’-end with [γ32P]ATP and polynucleotide kinase and purified in a 6% polyacrylamide gel as described (9).

In vitro structure probing and footprinting

For structure probing of NsiR1.1 and NsiR1.4, 0.1 pmol of labeled NsiR1.1 and NsiR1.4 RNAs in a 7 µL volume were denatured for 1 min at 95 °C and chilled on ice for 5 min. For the footprinting of NsiR1.1 on hetF 5’UTR, 0.1 pmol of labeled hetF 5’UTR RNA was mixed in 7 µL with water (-) or 1 pmol NsiR1.1 (+), denatured for 1 min at 95 °C and chilled on ice for 5 min. For the footprinting of NsiR1.1 or NsiR1.4 on alr3234 5’UTR, 0.1 pmol of labeled alr3234 (WT or MUT variant) 5’UTR RNAs were mixed in 7 µL with water (-) or 1 pmol NsiR1.1 or NsiR1.4 (+), denatured for 1 min at 95 °C and chilled on ice for 5 min. After denaturing and chilling steps, we added to all samples 1 µL of 1 mg/mL yeast RNA (Ambion AM7118) and 1 µL of 10x structure buffer (Ambion). The samples were incubated further for 15 min at 37 °C. RNase T1, RNase A and lead(II) acetate treatment were performed as described (9). Alkaline and RNase T1 G ladders were generated as described (9). All samples were run on 6% polyacrylamide, 7 M urea gels and bands visualized with a Cyclone Storage Phosphor System.

RNA isolation, Northern blot analysis and primer extension assays

Total RNA was isolated using hot phenol as described (10) with modifications (11). Samples for Northern blot hybridization of NsiR1 were separated in 10% urea-polyacrylamide gels as described (12) (7 µg of total RNA). Samples for Northern blot hybridization of longer

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transcripts (hetR or nifH) were separated in 1% agarose denaturing formaldehyde gels (10 µg of total RNA) and transferred to Hybond-N+ membrane (Amersham) with 20X SSC buffer.

Strand-specific 32P-labeled probes were prepared with Taq DNA polymerase using a PCR fragment as template and the oligonucleotides specified in SI Appendix, Table S3 in a reaction with [γ-32P]dCTP and one single oligonucleotide as primer (corresponding to the complementary strand of the sRNA or mRNA to be detected). PCR fragments for NsiR1.1 and NsiR1.4 probes were amplified with oligonucleotides 573 + 506 and pMBA77 (NsiR1.1) or pMBA78 (NsiR1.4) as templates. PCR fragments for hetR and nifH probes were amplified from genomic DNA using oligonucleotides hetR8 + hetR9 and 787 + 788 respectively. Hybridization to rnpB (13) or 5S rRNA probes was used as a loading and transfer control.

Primer extension of 5´ ends of hetF was performed as previously described (14) using 10 µg of total RNA and oligonucleotide 105 labeled with [γ-32P]ATP.

Heterocyst differentiation and commitment assay

Frequency of heterocysts was determined taking samples at different time points after nitrogen removal and staining with Alcian blue as described (15).The frequency of heterocysts was calculated as the number of cells stained by Alcian blue per at least 500 individual cells counted. All results are expressed as the average of 2 replicates ± SD. To assay heterocyst commitment, cells of the OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains were taken from nitrate-containing plates and grown in 100 mL of liquid BG110 + 4 mM ammonia for 72 h before being subjected to nitrogen deficiency. Nitrogen deficiency was induced by filtration and washing of the cells as described above. Washed cells were resuspended in 100 mL of nitrogen- free BG110 medium and incubated at 30ºC with shaking. At different times 2 mL were transferred to a glass tube, supplemented with 4 mM ammonia and further incubated for 24-36 h before heterocyst counting.

Nitrate and nitrite reductase assay

OE_C and OE_NsiR1.4 cultures were grown photoautotrophically at 30ºC and bubbled with an air/CO2 mixture (1% v/v) in BG110C + 6 mM NH4. Exponentially growing cells were harvested by filtration, washed with BG110C and resuspended in three different nitrogen sources. Aliquots of 20 mL at 4 µg chlorophyll/mL were incubated at 30ºC in BG110C (N2), BG11C

- + + (BG11 medium with 10 mM NaHCO3) (NO3 ) or BG110C + 6 mm NH4 (NH4 ) bubbled with an

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air/CO2 mixture (1% v/v). After 4 hours of incubation, cells were concentrated by centrifugation to a final concentration of 50 µg chlorophyll/mL. Nitrate reductase (16) and nitrite reductase (17) assays were performed with an amount of cells corresponding to 5 and 25 µg chlorophyll respectively. Dithionite-reduced methyl viologen was used as the reductant in cells made permeable with mixed alkyltrimethylammonium bromide.

Western blot analysis

For HetR Western blot analysis, crude extracts were prepared using glass beads. Cells from 25 mL cultures of OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 subjected to 18 h of nitrogen deficiency were harvested by filtration, washed with TE50/100 buffer (50 mM Tris pH 8, 100 mM EDTA) and resuspended in 500 µL of resuspension buffer (50 mM Tris-HCl pH 8.0, 2 mM 2-mercaptoethanol) containing a protease inhibitor cocktail (Roche). The cellular suspension was mixed with glass beads (SIGMA, 200 µm) in an Eppendorf tube and subjected to 7 cycles of 1 min vortexing plus 1 min of cooling on ice. Cell extract was separated from cell debris and unbroken cells by centrifugation (3 min at 3000 x g at 4 °C). The soluble fraction was obtained by centrifugation of the crude extract at 16.000 x g for 30 min at 4 °C. The protein concentration was determined by the Lowry procedure (18). 40 µg of proteins were fractionated on 10% SDS-PAGE. Antibodies against Nostoc sp. PCC 7120 HetR (19) and E. coli GroEL (Sigma- Aldrich) were used.

For Alr3234::GFP Western blot analysis, E.coli cells from stationary phase cultures were harvested and resuspended in SDS-PAGE loading buffer. Proteins were fractionated in 10% SDS-PAGE. Antibodies against GFP (Roche) and E. coli GroEL (Sigma-Aldrich) were used.

ECL Plus immunoblotting system (GE Healthcare) was used to detect the different primary antibodies using anti-rabbit (Sigma-Aldrich) or anti-mouse (Bio-Rad) horse-radish peroxidase conjugated secondary antibodies.

Reporter assay for in vivo verification of targets

We used the GFP reporter system described in (20) with the superfolder GFP (sfGFP) and plasmid pXG10-SF (21) for the experimental verification of targets in E.coli. The 5’UTR of hetF from the TSS at position –257 (4273166f) plus 60 nucleotides within its coding region was amplified using oligonucleotides 555 + 557. The GTG start codon was changed to ATG using

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overlapping PCRs and oligonucleotides 555, 556, 557 and 558. The information about TSS was taken from (22).

The whole intergenic region alr3233-alr3234 plus 60 nucleotides within the coding region of alr3234, containing the predicted NsiR1 interaction sequence, was amplified using oligonucleotides 553 + 870.

PCR products containing hetF::sfgfp and alr3234::sfgfp were digested with NsiI-NheI and cloned into NsiI-NheI-digested pXG10-SF vector, resulting in translational fusions of truncated HetF or Alr3234 to sfGFP (plasmids pMML2 and pMBA86, respectively, SI Appendix, Table S2).

To express NsiR1.1 and NsiR1.4 in E.coli, the genes were amplified from genomic DNA using primers 559 (5’ phosphorylated) + 561 or 560 (5’ phosphorylated) + 562, respectively. These PCR products were digested with XbaI and fused to a plasmid backbone that was amplified from pZE-12luc with primers PlacOB and PLlacOD and digested with XbaI, rendering pMML3 and pMML4 (SI Appendix, Table S2).

For the mutagenesis of 5’UTR of alr3234, a plasmid similar to pMBA86 was constructed using, in addition to the mentioned oligonucleotides, overlapping PCR with primers 871 + 872 containing the desired changes (pMBA87, SI Appendix, Table S2 and Table S3). The positions for the mutation of alr3234 5’UTR were selected based on hybridization energies predicted by IntaRNA (23).

Different combinations of plasmids were introduced in E. coli DH5α and GFP fluorescence measurements were done by flow cytometry or with a microplate reader (Varioskan) using liquid cultures from individual colonies, as previously described (24). Sequences of inserts from plasmids used in the heterologous reporter system are shown in SI Appendix, Table S5.

Fluorescence microscopy

Fluorescence of Nostoc sp. PCC 7120 filaments carrying plasmid pMBA92, pMBA93 and pMBA94 (SI Appendix, Table S6) growing on top of solidified nitrogen-free medium, were analyzed and quantified as described (25) using a Leica HCX PLAN-APO 63× 1.4 NA oil immersion objective attached to a Leica TCS SP2 laser-scanning confocal microscope. Samples were excited at 488 nm by an argon ion laser and the fluorescent emission was monitored by collection across windows of 500–538 nm (GFP) and 630–700 nm (cyanobacterial autofluorescence).

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Computational methods

To detect nsiR1 homologs in the upstream region of hetF, the HetF sequence of Nostoc sp. PCC 7120 was used in a blastp search in IMG (26) against 100 cyanobacterial genomes, that were those used in (27), excluding those corresponding to the fast-evolving Prochlorococcus clade (SI Appendix, Figure S1). We took 4 kb upstream from the 50 hetF homologs that were identified and used one of the identical repeats of nsiR1 in Nostoc sp. PCC 7120 as input in a blastn search to predict putative nsiR1 homologs in sequences upstream the predicted hetF homologs. Sequences of nsiR1 homologs that were not in the upstream region of hetF were taken from (11).The interaction between NsiR1.1 and NsiR1.4 with alr3234 mRNA was analyzed using intaRNA (23).

Quantification and statistical analysis

The students’ t test was used to determine statistical significance. Number of biological samples can be found in the figure legends.

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Figure S1. Co-occurrence of nsiR1 and hetF. Genomes are arranged and color-coded on the left as in (27) as follows: filamentous non-heterocystous in green, heterocystous ramified in blue, heterocystous non-ramified in pink, baeocystous in orange and unicellular in black. Presence or absence of a hetF homolog is indicated. Presence or absence of homologs of nsiR1, number of putative nsiR1 copies and genomic location of the nsiR1 arrays upstream to hetF (light green) or in a different genomic region (yellow) are also indicated. Nostoc sp. PCC 7120 is highlighted in red.

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Figure S2. Genomic regions of nsiR1 repeats in 40 genomes. nsiR1 repeats are indicated by small arrows with solid colors (red for identical repeats, light blue for repeats with sequence divergence and green for repeats that might be antisense to hetF 5’-UTR). Genes are indicated by arrows with faded colors. Homologous genes are shown in the same color. hetF homologs are shown by dark grey arrows on the right side of the regions. The TSS of hetF in Nostoc sp. PCC 7120 is indicated by a bent arrow. The coherent clade of heterocyst-forming cyanobacteria is highlighted by a red frame. Nostoc sp. PCC 7120 is indicated in red.

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Figure S3. Structure probing of NsiR1.1 and NsiR1.4. (A) Secondary structure model of NsiR1.1 and NsiR1.4. Black and white triangles indicate the positions sensitive to hydrolysis by RNase T1 or lead(II) acetate (Pb), respectively. (B) 5'-end labelled NsiR1.1 (left) or NsiR1.4 (right) were incubated with RNase T1 or lead(II) acetate and the resulting fragments analyzed on an 8% polyacrylamide sequencing gel. C, untreated control; OH, alkaline ladder; T1s, RNase T1 ladder. Nucleotide positions of NsiR1.1 or NsiR1.4 are shown on the left.

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Figure S4. Accumulation of hetR transcripts in OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains. (A) hetR expression. Northern blots with RNA isolated from the indicated strains at different time points (indicated in hours) after removal of combined nitrogen were hybridized with probes for hetR (top panels) and rnpB (bottom panels), used as a loading control. Membranes are the same used in Fig 4D. Sizes (in nucleotides) are indicated on the left. (B) Quantification of hetR expression. The data are normalized to the amount of rnpB.

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Figure S5. Heterocyst differentiation in strains with altered amounts of NsiR1. Bright field images of filaments of + OE_C, OE_NsiR1.1, OE_NsiR1.4 and OE_as_NsiR1 strains growing in liquid media containing ammonium (NH4 ) or growing in liquid media without combined nitrogen 24 h after nitrogen removal (N2). Filaments with enlarged irregular cells are pointed by black arrows. Scale bars, 25 µm.

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Table S1. Strains

Strain Description Reference Escherichia coli DH5α Used for routine transformation (28) Nostoc sp. PCC 7120 Wild type Pasteur Culture Collection 216 Non-functional HetR. S179N mutation. (1) OE_C SmRSpR, pMBA51 inserted in plasmid alpha. T1 terminator (5) of E. coli rrnB gene expressed constitutively from the trc promoter. OE_NsiR1.1 SmRSpR, pMBA77 inserted in plasmid alpha. NsiR1.1 This work transcribed constitutively from the trc promoter. OE_NsiR1.4 SmRSpR, pMBA78 inserted in plasmid alpha. NsiR1.4 This work expressed constitutively from trc promoter. OE_as_NsiR1 SmRSpR, pMBA42 inserted in plasmid alpha. Antisense to This work NsiR1 transcribed constitutively from the trc promoter.

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Table S2. Plasmids

Name Description Reference pJV300 Control plasmid expresses a ~50 nt transcript derived from the T1 (29) terminator of E.coli rrnB gene. pRL278 NmR, shuttle vector for conjugation in Nostoc with a Neomycin resistance (7) cassette. pMBA20 ApRSmRSpR, plasmid for the overexpression of transcripts from the rnpB (5) promoter and followed by the T1 transcriptional terminator of E. coli rrnB gene. pMBA37 ApRSmRSpR, plasmid for the overexpression of transcripts from the trc (5) promoter and followed by the T1 terminator of E. coli rrnB gene, used as transcriptional terminator. pMBA51 ApRSmRSpR, control plasmid, expresses a 56 nt transcript derived from (5) the T1 terminator of E. coli rrnB gene from the trc promoter. pXG0 CmR, control plasmid without GFP (20) pXG10-SF CmR, plasmid for construction of translational sfGFP fusions. (21) R pZE12-luc Ap , plasmid used for the expression of NsiR1 under control of the PLlac-O (30) promoter. pMBA42 ApRSmRSpR, plasmid based on pMBA37 for the overexpression of an This work antisense to NsiR1.1. pMBA77 ApRSmRSpR, plasmid based on pMBA37 for the overexpression of NsiR1.1. This work pMBA78 ApRSmRSpR, plasmid based on pMBA37 for the overexpression of NsiR1.4. This work pMBA86 CmR, sfGFP reporter plasmid based on pXG10-SF containing the alr3234 This work 5’UTR plus sequences encoding the first 20 amino acids of Alr3234. pMBA87 CmR, same as pMBA86 but with T4G and G9A substitutions in the alr3234 This work 5’UTR (alr3234 MUT). pMBA91 ApRNmR, Kanamycin/ Neomycin resistance version of pMBA20. This work pMBA92 ApRNmR, plasmid based on pMBA91 expressing from rnpB promoter the This work gene gfpmut2 plus its 5’UTR. pMBA93 ApRNmR, plasmid based on pMBA91 expressing from the rnpB promoter This work a translational fusion of hetF 5’UTR plus sequences encoding the first 20 amino acids of HetF fused to GFP. pMBA94 ApRNmR, plasmid based on pMBA91 expressing from rnpB promoter a This work translational fusion of alr3234 5’UTR plus sequences encoding the first 20 amino acids of Alr3234 fused to GFP. pMML2 CmR, sfGFP reporter plasmid based on pXG10-SF containing the hetF This work 5’UTR plus sequences encoding the first 20 amino acids of HetF (with mutation in start codon GTG -> ATG). pMML3 ApR, plasmid based on pZE12-luc expressing NsiR1.1. This work pMML4 ApR, plasmid based on pZE12-luc expressing NsiR1.4. This work

203

Table S3. Oligonucleotides

Name Sequence (5’-3’) Used for 105 CGCCAAAGTTTCAATTCGATTTCC Primer extension of hetF 189 CGCACTGACCGAATTCATTAA Plasmid backbone (PLlacOB) amplification from pZE12- 190 GTGCTCAGTATCTTGTTATCCG luc (PLlacOD) 506 GTTTTGAGCTCAGGTCTAGGGCGGCGGATTT Probes for nsiR1.1 and nsiR1.4 512 TTTTATCTCGAGGTAGATGCACCCTGATAACTAACTC Cloning of as_nsiR1 fused to CCCTAGCTGGCTAACACCGACTGGGGGCTTTTTATGC the trc promoter in pMBA37 ATGCTCCAGTGTAATC 553 GTTTTATGCATACAATCATTAAAAATGTGGTGCAG Cloning of alr3234 5’UTR in 870 GTTTTGCTAGCTACTTGCTCATTATTTATTTTC pxG10-SF 555 GTTTTATGCATAAGTATATGCTAGTTATAGAAAAAAC Cloning of hetF 5’UTR with CCCC mutation in start codon 556 GAAATTCCTGGGACATAACTTCTCCTATC (GTG->ATG) in pxG10-SF 557 GTTTTGCTAGCCCGCACCAAGTAGTCATTCTGCC 558 GATAGGAGAAGTTATGTCCCAGGAATTTC 559 5’P-GGTAGATGCACCTTGATCTTTAA Cloning of nsiR1.1 in pZE12- 561 GTTTTTTCTAGACCCAAGACAGTAGAATAATCTAAG luc 560 5’P-GGTAGATGCACCCTGATAACTA Cloning of nsiR1.4 in pZE12- 562 GTTTTTTCTAGACGGATGAATCTGCACAATGATG luc 573 GTTTTATCGATGCGCCGACATCATAACGGTT Cloning of the trc promoter 574 GTTTTATGCATTCCACACATTATACGAGCCG 757 GTTTTATCTCGAGAAAAAACCCCCAGTGGCTATCAAC Cloning of nsiR1.1 fused to CAACTAGGGGAGTTAAAGATCAAGGTGCATCTACCAT the trc promoter in pMBA37 GCATTCCACACATTATACG 758 GTTTTATCTCGAGAAAAAGCCCCCAGTCGGTGTTAGC Cloning of nsiR1.4 fused to CAGCTAGGGGAGTTAGTTATCAGGGTGCATCTACCAT the trc promoter in pMBA37 GCATTCCACACATTATACG 787 GTACTGCAAGGGGCGTGGCT Probe for nifH 788 CCTATTGGTAGCTTCTGCGGG 831 TAATACGACTCACTATAGGGAAGTATATGCTAGTTAT In vitro transcription from AGAAAAAACCCCC the T7 promoter of hetF 5’UTR 833 TAATACGACTCACTATAGGGTAGATGCACCTTGATCT In vitro transcription from TTAACTCCCCTAGTTGGTTG the T7 promoter of NsiR1.1 834 AAAAAACCCCCAGTGGCTATCAACCAACTAGGGGAGT TAAAGATC 835 TAATACGACTCACTATAGGGTAGATGCACCCTGATAA In vitro transcription from CTAACTCCCCTAGCTGGCTAAC the T7 promoter of NsiR1.4 836 AAAAAAGCCCCCAGTCGGTGTTAGCCAGCTAGGGGAG TTAG 871 AATGTCTTCTTGATCCATAACTTCCCCTGAT Mutagenesis of alr3234 872 AAGTTATGGATCAAGAAGACATTTACAATTC 5’UTR 873 CCGCACCAAGTAGTCATTCTG In vitro transcription from the T7 promoter of hetF 5’UTR 878 TAATACGACTCACTATAGGGACAATCATTAAAAATGT In vitro transcription from GGTGCAG the T7 promoter of alr3234 879 TACTTGCTCATTATTTATTTTCTTAACG 5’UTR. hetR8 GCGCTCTGGTGGCACATCG Probe for hetR hetR9 CATGAGGACGAGCGGGCATA

204

Name Sequence (5’-3’) Used for 892 GTTTTGGATCCCTGCATCCCTTAACTTACTTAT Cloning of Nm resistance 893 GTCTAGAGGATCCCCGGTGGG cassette in pMBA20 894 GTTTTCTCGAGGCATGCCTGCAGGTCTGGACAT Cloning of translational fusions to GFP in pMBA91 895 CTCCTTTACTCCGCACCAAGTAGTCATTCTGC Cloning of hetF 5’UTR fused 896 TACTTGGTGCGGAGTAAAGGAGAAGAACTTTTC to GFP 897 TCTCCTTTACTTACTTGCTCATTATTTATTTTC Cloning of alr3234 5’UTR 898 ATGAGCAAGTAAGTAAAGGAGAAGAACTTTTC fused to GFP 899 GTTTTATGCATGCGAAAGCTTGCATGCCTGCAGG Cloning of gfpmut2 5’UTR

Restriction sites used for cloning (underlined), point mutations made in hetF start codon or alr3234 5’UTR (red) and 5’ monophosphate (5’P) are indicated.

205

Table S4. RNAs used for in vitro footprinting assays.

Name Sequence (5’-3’) NsiR1.1 gGGUAGAUGCACCUUGAUCUUUAACUCCCCUAGUUGGUUGAUAGCCACUGGGGG UUUUUU NsiR1.4 gGGUAGAUGCACCCUGAUAACUAACUCCCCUAGCUGGCUAACACCGACUGGGGG CUUUUUU hetF 5’UTR gggAAGUAUAUGCUAGUUAUAGAAAAAACCCCCAGUGGCUAUCAACCAACUAGG GGAGUUAAAGAUCAAGGUGCAUCUACCGUUAAACUGUUCUGGGCAGAUGCUAAC CAAUCCGGAUAAAGUUGUAGAUGUAGAACCAGCAAAACCUUUGAGUGGAGAAAC AUUCAAUGGCUUGCUGCAUCUAGAUUUACAGAAAAAAUGGAAAUCGAAUUGAAA CUUUGGCGUUUCAAGCUAGAUUUAGGAGGCAGAUAGGAGAAGUUGUGUCCCAGG AAUUUCACAUUUCUGUAACCCCAGUAGGGCAGAAUGACUACUUGGUGCGGUUCU GUAACCCCAGUAGGGCAGAAUGACUACUUGGUGCGG alr3234 5’UTR gggACAAUCAUUAAAAAUGUGGUGCAGACAUAUACCCAAACUUUAGAAACAUCA (WT) AUUUACUCUUAGUGGUAAUGGGUAAUAUAGCUGUUGACAGUUAACUGUUGACAG UUGACUGUUGACAGAUUUGAAAGUCUUGUUUUAUCAGGGUUUGAUGUUAGCUGG AUGUUUUAAUCUAUCUGGCUACAGCUAUAAAUGUAAUUCUCUUCUCCAUCACCA AUUACCAAUUACCAAUUACCAAUUUUCAAACCAUCAAAUAAAAUCAUCAGGGGA AGUUAUGUAUCAGGAAGACAUUUACAAUUCACAGAACGUUAAGAAAAUAAAUAA UGAGCAAGUA alr3324 5’UTR gggACAAUCAUUAAAAAUGUGGUGCAGACAUAUACCCAAACUUUAGAAACAUCA (MUT) AUUUACUCUUAGUGGUAAUGGGUAAUAUAGCUGUUGACAGUUAACUGUUGACAG UUGACUGUUGACAGAUUUGAAAGUCUUGUUUUAUCAGGGUUUGAUGUUAGCUGG AUGUUUUAAUCUAUCUGGCUACAGCUAUAAAUGUAAUUCUCUUCUCCAUCACCA AUUACCAAUUACCAAUUACCAAUUUUCAAACCAUCAAAUAAAAUCAUCAGGGGA AGUUAUGGAUCAAGAAGACAUUUACAAUUCACAGAACGUUAAGAAAAUAAAUAA UGAGCAAGUA

Nostoc sequences are capitalized, guanosines added for T7 transcription efficiency are in lower case. Predicted interaction sites of hetF 5’UTR and the two versions of alr3234 5’UTR with different versions of NsiR1 are highlighted in yellow. Start codons are shown in bold and underlined. Mutations introduced in alr3234 5’UTR MUT version are indicated in red.

206

Table S5. Sequences of inserts in plasmids used for verification of sRNA-mRNA interactions in E. coli.

Plasmid Sequence Description

pMML2 atgcatAAGTATATGCTAGTTATAGAAAAAACCCCCAGTGGCTATC hetF AACCAACTAGGGGAGTTAAAGATCAAGGTGCATCTACCGTTAAACT 5’UTR GTTCTGGGCAGATGCTAACCAATCCGGATAAAGTTGTAGATGTAGA ACCAGCAAAACCTTTGAGTGGAGAAACATTCAATGGCTTGCTGCAT CTAGATTTACAGAAAAAATGGAAATCGAATTGAAACTTTGGCGTTT CAAGCTAGATTTAGGAGGCAGATAGGAGAAGTTATGTCCCAGGAAT TTCACATTTCTGTAACCCCAGTAGGGCAGAATGACTACTTGGTGCG Ggctagc pMBA86 atgcatACAATCATTAAAAATGTGGTGCAGACATATACCCAAACTT alr3234 (WT) TAGAAACATCAATTTACTCTTAGTGGTAATGGGTAATATAGCTGTT 5’UTR GACAGTTAACTGTTGACAGTTGACTGTTGACAGATTTGAAAGTCTT GTTTTATCAGGGTTTGATGTTAGCTGGATGTTTTAATCTATCTGGC TACAGCTATAAATGTAATTCTCTTCTCCATCACCAATTACCAATTA CCAATTACCAATTTTCAAACCATCAAATAAAATCATCAGGGGAAGT TATGTATCAGGAAGACATTTACAATTCACAGAACGTTAAGAAAATA AATAATGAGCAAGTAgctagc pMBA87 atgcatACAATCATTAAAAATGTGGTGCAGACATATACCCAAACTT alr3234 (MUT) TAGAAACATCAATTTACTCTTAGTGGTAATGGGTAATATAGCTGTT 5’UTR GACAGTTAACTGTTGACAGTTGACTGTTGACAGATTTGAAAGTCTT GTTTTATCAGGGTTTGATGTTAGCTGGATGTTTTAATCTATCTGGC TACAGCTATAAATGTAATTCTCTTCTCCATCACCAATTACCAATTA CCAATTACCAATTTTCAAACCATCAAATAAAATCATCAGGGGAAGT TATGGATCAAGAAGACATTTACAATTCACAGAACGTTAAGAAAATA AATAATGAGCAAGTAgctagc pMML3 GGTAGATGCACCTTGATCTTTAACTCCCCTAGTTGGTTGATAGCCA NsiR1.1 CTGGGGGTTTTTTCTATAACTAGCATATACTTAGATTATTCTACTG TCTTGGGtctaga pMML4 GGTAGATGCACCCTGATAACTAACTCCCCTAGCTGGCTAACACCGA NsiR1.4 CTGGGGGCTTTTTTATGTTCATAAATAACCAGCATCATTGTGCAGA TTCATCCGTCAGATCTAGAtctaga

Nostoc sequences are capitalized, with black letters corresponding to 5’UTR and blue letters corresponding to coding sequences, respectively. NsiI and NheI sites that were used for cloning are highlighted in magenta and yellow, respectively. Nucleotide changes with respect to wild type sequences are indicated in red. Grey shadowed letters indicate the NsiR1.1 and NsiR1.4 sequences. XbaI restriction sites used for cloning are highlighted in blue. Start codons are shown in bold and underlined.

207

Table S6. Sequences of inserts in plasmids expressing GFP translational fusions in Nostoc.

Plasmid Sequence Description

pMBA92 GTCCCTTCACTATCAAAAAACTACCTTGGACTTATGCCCTACCCTG gfpmut2 plus its GAATAAAAAGAAATAAGCAAAACAGACACAAGACACCAACGAAGAT original 5’UTR TACACTGGAGCatgcatgcgaaagcttgcatgcctgcaggtcgact downstream the ctagaggatcctctagatttaagaaggagatatacatatgagtaaa rnpB promoter ggagaagaacttttcactggagttgtcccaattcttgttgaattag atggtgatgttaatgggcacaaattttctgtcagtggagagggtga aggtgatgcaacatacggaaaacttacccttaaatttatttgcact actggaaaactacctgttccatggccaacacttgtcactactttcg cgtatggtcttcaatgctttgcgagatacccagatcatatgaaaca gcatgactttttcaagagtgccatgcccgaaggttatgtacaggaa agaactatatttttcaaagatgacgggaactacaagacacgtgctg aagtcaagtttgaaggtgatacccttgttaatagaatcgagttaaa aggtattgattttaaagaagatggaaacattcttggacacaaattg gaatacaactataactcacacaatgtatacatcatggcagacaaac aaaagaatggaatcaaagttaacttcaaaattagacacaacattga agatggaagcgttcaactagcagaccattatcaacaaaatactcca attggcgatggccctgtccttttaccagacaaccattacctgtcca cacaatctgccctttcgaaagatcccaacgaaaagagagaccacat ggtccttcttgagtttgtaacagctgctgggattacacatggcatg gatgaactatacaaataaatgtccagacctgcaggcatgcctcgag pMBA93 GTCCCTTCACTATCAAAAAACTACCTTGGACTTATGCCCTACCCTG Translational GAATAAAAAGAAATAAGCAAAACAGACACAAGACACCAACGAAGAT fusion of hetF TACACTGGAGCatgcatAAGTATATGCTAGTTATAGAAAAAACCCC 5’UTR plus CAGTGGCTATCAACCAACTAGGGGAGTTAAAGATCAAGGTGCATCT sequences ACCGTTAAACTGTTCTGGGCAGATGCTAACCAATCCGGATAAAGTT encoding the GTAGATGTAGAACCAGCAAAACCTTTGAGTGGAGAAACATTCAATG first 20 amino GCTTGCTGCATCTAGATTTACAGAAAAAATGGAAATCGAATTGAAA CTTTGGCGTTTCAAGCTAGATTTAGGAGGCAGATAGGAGAAGTTGT acids of HetF GTCCCAGGAATTTCACATTTCTGTAACCCCAGTAGGGCAGAATGAC fused to GFP, TACTTGGTGCGGagtaaaggagaagaacttttcactggagttgtcc downstream the caattcttgttgaattagatggtgatgttaatgggcacaaattttc rnpB promoter tgtcagtggagagggtgaaggtgatgcaacatacggaaaacttacc cttaaatttatttgcactactggaaaactacctgttccatggccaa cacttgtcactactttcgcgtatggtcttcaatgctttgcgagata cccagatcatatgaaacagcatgactttttcaagagtgccatgccc gaaggttatgtacaggaaagaactatatttttcaaagatgacggga actacaagacacgtgctgaagtcaagtttgaaggtgatacccttgt taatagaatcgagttaaaaggtattgattttaaagaagatggaaac attcttggacacaaattggaatacaactataactcacacaatgtat acatcatggcagacaaacaaaagaatggaatcaaagttaacttcaa aattagacacaacattgaagatggaagcgttcaactagcagaccat tatcaacaaaatactccaattggcgatggccctgtccttttaccag acaaccattacctgtccacacaatctgccctttcgaaagatcccaa cgaaaagagagaccacatggtccttcttgagtttgtaacagctgct gggattacacatggcatggatgaactatacaaataaatgtccagac ctgcaggcatgcctcgag pMBA94 GTCCCTTCACTATCAAAAAACTACCTTGGACTTATGCCCTACCCTG Translational GAATAAAAAGAAATAAGCAAAACAGACACAAGACACCAACGAAGAT fusion of TACACTGGAGCatgcatACAATCATTAAAAATGTGGTGCAGACATA alr3234 5’UTR TACCCAAACTTTAGAAACATCAATTTACTCTTAGTGGTAATGGGTA plus sequences ATATAGCTGTTGACAGTTAACTGTTGACAGTTGACTGTTGACAGAT encoding the TTGAAAGTCTTGTTTTATCAGGGTTTGATGTTAGCTGGATGTTTTA first 20 amino ATCTATCTGGCTACAGCTATAAATGTAATTCTCTTCTCCATCACCA ATTACCAATTACCAATTACCAATTTTCAAACCATCAAATAAAATCA acids of Alr3234 TCAGGGGAAGTTATGTATCAGGAAGACATTTACAATTCACAGAACG fused to GFP, TTAAGAAAATAAATAATGAGCAAGTAagtaaaggagaagaactttt

208

cactggagttgtcccaattcttgttgaattagatggtgatgttaat downstream the gggcacaaattttctgtcagtggagagggtgaaggtgatgcaacat rnpB promoter acggaaaacttacccttaaatttatttgcactactggaaaactacc tgttccatggccaacacttgtcactactttcgcgtatggtcttcaa tgctttgcgagatacccagatcatatgaaacagcatgactttttca agagtgccatgcccgaaggttatgtacaggaaagaactatattttt caaagatgacgggaactacaagacacgtgctgaagtcaagtttgaa ggtgatacccttgttaatagaatcgagttaaaaggtattgatttta aagaagatggaaacattcttggacacaaattggaatacaactataa ctcacacaatgtatacatcatggcagacaaacaaaagaatggaatc aaagttaacttcaaaattagacacaacattgaagatggaagcgttc aactagcagaccattatcaacaaaatactccaattggcgatggccc tgtccttttaccagacaaccattacctgtccacacaatctgccctt tcgaaagatcccaacgaaaagagagaccacatggtccttcttgagt ttgtaacagctgctgggattacacatggcatggatgaactatacaa ataaatgtccagacctgcaggcatgcctcgag

Nostoc sequences are capitalized, with black letters corresponding to 5’UTR, blue letters corresponding to coding sequences and orange letters corresponding to the rnpB promoter, respectively. NsiI and XhoI sites that were used for cloning are highlighted in magenta and red, respectively. Start codons are shown in bold and underlined.

209

Supplementary references

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15. E. Olmedo-Verd, E. Flores, A. Herrero, A. M. Muro-Pastor, HetR-dependent and - independent expression of heterocyst-related genes in an Anabaena strain overproducing the NtcA transcription factor. J. Bacteriol. 187, 1985-1991 (2005).

16. A. Herrero, E. Flores, M. G. Guerrero, Regulation of nitrate reductase cellular levels in the cyanobacteria Anabaena variabilis and Synechocystis sp. FEMS Microbiol. Lett. 26, 21-25 (1985).

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19. A. Valladares, E. Flores, A. Herrero, The heterocyst differentiation transcriptional regulator HetR of the filamentous cyanobacterium Anabaena forms tetramers and can be regulated by phosphorylation. Mol. Microbiol. 99, 808-819 (2016).

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21. C. P. Corcoran et al., Superfolder GFP reporters validate diverse new mRNA targets of the classic porin regulator, MicF RNA. Mol. Microbiol. 84, 428-445 (2012).

22. J. Mitschke, A. Vioque, F. Haas, W. R. Hess, A. M. Muro-Pastor, Dynamics of transcriptional start site selection during nitrogen stress-induced cell differentiation in Anabaena sp. PCC 7120. Proc. Natl. Acad. Sci. U.S.A 108, 20130-20135 (2011).

23. M. Mann, P. R. Wright, R. Backofen, IntaRNA 2.0: enhanced and customizable prediction of RNA-RNA interactions. Nucleic Acids Res. 45, W435-W439 (2017).

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25. A. M. Muro-Pastor, The heterocyst-specific NsiR1 small RNA is an early marker of cell differentiation in cyanobacterial filaments. mBio 5, e01079-01014 (2014).

26. V. M. Markowitz et al., IMG 4 version of the integrated microbial genomes comparative analysis system. Nucleic Acids Res. 42, D560-D567 (2014).

27. P. M. Shih et al., Improving the coverage of the cyanobacterial phylum using diversity- driven genome sequencing. Proc. Natl. Acad. Sci. U.S.A 110, 1053-1058 (2013).

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4. GENERAL DISCUSSION

General discussion

4. GENERAL DISCUSSION

RNA regulators play an important role in the regulation of gene expression in bacteria (Wade and Grainger, 2014). In the last decades, the increase in the number of available genomes, the design of specific computational software and the increase of available “omic” data have allowed the detection and characterization of hundreds of new RNA regulators (Wagner and Romby, 2015).

In cyanobacteria, some computational approaches have been carried out to detect sRNAs (Axmann et al., 2005; Voss et al., 2009), showing that a high number of RNA regulators may be encoded in cyanobacterial genomes. Some cyanobacterial asRNAs and sRNAs are involved in the regulation of photosynthesis (Wilde and Hihara, 2016), a central process in a photosynthetic organism. In addition, some sRNAs seem to be involved in the regulation of adaptation to several stresses, such as iron limitation (Georg et al., 2017) or nitrogen limitation (Klähn et al., 2015).

Most of the previously studied cyanobacterial sRNAs and asRNAs were found in the model unicellular cyanobacterium Synechocystis sp. PCC 6803. However, in our group we are interested in the regulation exerted by this type of molecules in Nostoc sp. PCC 7120, a multicellular microorganism that can undergo a cellular differentiation process. So, the objectives of this Thesis have been focused in the identification of new sRNAs and asRNAs and the characterization of some of them that could affect the regulation of heterocyst differentiation.

4.1 A computational approach for the identification of conserved sRNAs in heterocyst- forming cyanobacteria.

The increasing number of available genomes allows the design of computational approaches to the prediction of possible sRNAs. The first predictions carried out in E. coli were based on the conservation of sequences (or structure motifs) located in intergenic regions and flanked by transcription initiation signals and putative Rho-independent terminators (Argaman et al., 2001; Rivas et al., 2001; Wassarman et al., 2001; Chen et al., 2002). Some computational predictions were also carried out in cyanobacteria (Axmann et al., 2005; Voss et al., 2009). However, whereas the prediction of Rho-independent terminators is quite accurate, the prediction of promoter signals is not, producing many false positives. In our approach, we

215 General discussion

overcame this problem by using an experimental dataset (dRNAseq) that contained all the experimentally determined TSS in the genome of our model organism (Mitschke et al., 2011b). Therefore, we selected only sequences flanked by a previously determined TSS and a predicted Rho-independent terminator (Chapter I, Figure 1).

Our computational approach predicted 327 sRNAs that could be transcribed from the genome of Nostoc sp. PCC 7120, a number of sRNAs that is in the range (hundreds of sRNAs) predicted in the smaller genome of E. coli (Vogel and Sharma, 2005). We experimentally verified the expression of 14 previously undescribed sRNAs by Northern blot and in every case a transcript with the predicted size was identified. In addition, some of the predicted sRNAs had been previously identified in several studies, including Yfr1 and Yfr2 (Axmann et al., 2005), SyR22/IsaR1 (Georg et al., 2017), NsiR1 (Ionescu et al., 2010), NsiR3 (Mitschke et al., 2011b) and NsrR1 (Álvarez-Escribano et al., 2018). These validations confirm the good performance of our computational prediction.

The experimental dataset containing all TSS was obtained from samples corresponding to standard growth conditions or 8 hours of nitrogen deficiency. Therefore, transcripts induced under other specific stress conditions, and their TSS, are underrepresented in the dataset. The same argument holds true for sRNAs that are too small to be effectively included in the RNAseq data. For these reasons, in the predictive algorithm we also included TSS appearing in the RNAseq dataset with small numbers of sequencing reads. In fact, we tested expression of one of the predicted sRNAs associated to a TSS with only eight reads and its expression and size were confirmed by Northern blot (Chapter I, Figure 4), again validating the predictive approach. Recently developed software for the prediction of sRNA homologs is based on the conservation of primary sequences but also in the conservation of structural features (Eggenhofer et al., 2016; Lott et al., 2018). However, our results pointed to the importance of primary sequence conservation in the prediction of sRNAs with a small size (around 60 nucleotides), because this small size prevents folding of complex secondary structures.

Some of the predicted sRNAs were encoded in several copies in the genome, forming what we called “families”. The presence of sRNAs with multiples copies has been reported in the genome of several bacteria (Wilderman et al., 2004; Lenz et al., 2005; Guillier and Gottesman, 2008; Billenkamp et al., 2015). Among the predicted sRNAs encoded by multiple copies was NsiR1, a sRNA specifically induced in heterocysts (Ionescu et al., 2010). The functional divergence of the multiple copies of this sRNA and its implication in the regulation of heterocyst differentiation is the subject of Chapter IV.

216 General discussion

The predicted sRNAs showed different degrees of phylogenetic conservation (Chapter I, Figure 2). As expected, homologs to the predicted sRNAs were more abundant among heterocystous cyanobacteria because the prediction was carried out using a genome of a heterocystous strain (Nostoc sp. PCC 7120). Some sRNAs, as Yfr1 or Yfr2, were conserved in many cyanobacterial genomes pointing to a general function in the physiology of this group of organisms. Our results showed a strong conservation of a CACUCCUCACACC motif near the 5’ of Yfr1 (Chapter I, Figure 3) that may participate in the interaction between Yfr1 and its mRNA targets. This result had been previously reported in a phylogenetic analysis carried out with less cyanobacterial genomes (Voss et al., 2007). The functional characterization of Yfr1 is the subject of Chapter II.

We were particularly interested in sRNAs that were only conserved in the genomes of heterocystous cyanobacteria (Chapter I, Table 1). Our hypothesis was that if a sRNA was only conserved in heterocystous strains, it could be involved in some specific metabolic or physiological process performed by these cyanobacteria, including heterocyst differentiation. We verified the expression of several sRNAs that were induced after nitrogen deprivation and were only conserved in heterocystous cyanobacteria (Chapter I, Figure 5). The expression of some of them (NsiR8 and NsiR9) was HetR-dependent and consequently they could be specifically involved in the regulation of heterocyst differentiation. In fact, cell-specific transcription of NsiR8 was confirmed by using a fusion to gfp (Chapter I, Figure 6).

4.2 Yfr1, a widely conserved sRNA, regulates the integrity of cell wall and its remodeling during heterocyst differentiation.

Yfr1 was first identified in Prochlorococcus strains (Axmann et al., 2005). A later study (Voss et al., 2007) and our computational approach (Brenes-Álvarez et al., 2016) showed that Yfr1 is conserved in all analysed cyanobacterial genomes. The conservation of this sRNA in organisms with such diverse morphologies and ecological niches suggests an important function in the cyanobacterial physiology.

A prediction of possible targets was carried out using CopraRNA (Wright et al., 2013; Wright et al., 2014) and several homologs of Yfr1 identified by our computational approach in heterocystous cyanobacteria. The results of the target prediction showed an enrichment in genes encoding outer membrane proteins and enzymes involved in the synthesis and remodeling of peptidoglycan (Chapter II, Table S1).

217 General discussion

Eleven predicted targets were validated using a heterologous reporter system in E. coli (Corcoran et al., 2012). The expression of ten of them was downregulated in the presence of Yfr1 (Chapter II, Figure 1). The interaction site of two downregulated genes, murC and mraY, was mapped (Chapter II, Figure 2), confirming the prediction of interaction sites. The predicted interaction sites for these ten downregulated genes were located near the start codon, suggesting a negative regulation based on the interference of ribosome access (Chapter II, Figure S2).

Strains of Nostoc sp. PCC 7120 with altered levels of Yfr1 were generated. We overexpressed Yfr1 (OE_Yfr1 strain) and, following a strategy similar to that in Dühring et al., 2006, we tried to deplete the levels of endogenous Yfr1 through the overexpression of an antisense transcript that would act as a sponge interacting with endogenous Yfr1 (OE_as_Yfr1 strain). The expression of Yfr1 and two of the downregulated genes, murC and mraY, was tested in the Nostoc strains (Chapter II, Figure 3). The amount of full-length mRNA of these two mRNA was increased in the strain with a lower level of Yfr1 (OE_as_Yfr1), confirming the negative regulation exerted by Yfr1 on these targets previously validated in E. coli. These results suggested that the block of translation initiation by the interaction of Yfr1 with these two target mRNAs may favour its degradation by different RNases.

The Nostoc strains with altered levels of Yfr1 grew poorly in media with vancomycin, an antibiotic that interacts with nascent peptidoglycan (Chapter II, Figure 4). This result suggested that the peptidoglycan biosynthesis was altered in these strains, consistent with a downregulation of murC and mraY by Yfr1. The strain that overexpressed Yfr1 (OE_Yfr1) was also sensitive to SDS or erythromycin (Chapter II, Figure 4), two harmful compounds that usually cannot pass the envelopes of cyanobacteria, suggesting additional alterations of the integrity of the envelopes in this strain. In this case, the disrupted envelope of OE_Yfr1 could be associated to the verified interaction of Yf1 with alr2269 (Omp85), a protein involved in outer membrane protein biogenesis (Nicolaisen et al., 2009b). In addition, the Nostoc strains with altered levels of Yfr1 showed alterations in the distribution and completion of septa (Chapter II, Figure 4). This unusual distribution of septa could be associated to the regulation exerted by Yfr1 on the expression of genes related to peptidoglycan biosynthesis (murC and mraY) and remodeling (amiC2), but also to the regulation exerted on conR expression (Chapter II, Figure 1), a gene encoding a protein of the LytR-CpsA-Psr superfamily involved in septum formation (Mella- Herrera et al., 2011). In addition, the strain that overexpressed Yfr1 was unable to grow in media without combined nitrogen due to disruption of the filaments at the connections between vegetative cells and heterocysts (Chapter II, Figure 5), which is in concordance with regulation

218 General discussion exerted by Yfr1 on genes that are essential for diazotrophic growth, such as murC, amiC2 or conR (Zhu et al., 2001; Mella-Herrera et al., 2011; Videau et al., 2016b; Bornikoel et al., 2017). Thus, our results highlight the importance of the regulation of peptidoglycan synthesis and remodeling during heterocyst formation.

Unlike most previously described cyanobacterial sRNAs, such as NsiR1 (Ionescu et al., 2010), NsrR1 (Álvarez-Escribano et al., 2018), NsiR4 (Klähn et al., 2015), IsaR1 (Georg et al., 2017) or PsrR1 (Georg et al., 2014) that change their expression under certain stress conditions, Yfr1 is transcribed constitutively, which is unusual for regulatory RNAs. This observation points to a putative regulation of Yfr1 activity based on controlled sequestration exerted by another RNA or by an RNA-binding protein. In Prochlorococcus, a recent report (Lambrecht et al., 2019) has shown that Yfr2, a sRNA induced under several stresses, interacts with Yfr1, preventing the interaction between Yfr1 and its target mRNAs. In this type of regulatory circuit, the regulatory capacity of a sRNA that is expressed constitutively depends on the presence or absence of an interacting sRNA induced under different stresses (Chapter II, Figure 6).

4.3 A co-expression network to dissect the complex transcriptome of Nostoc sp. PCC 7120 during heterocyst differentiation.

In heterocyst-forming cyanobacteria, two responses take place simultaneously under nitrogen deprivation; a general response under control of NtcA and the differentiation of heterocysts that additionally depends on the master regulator HetR. In this work, we tried to dissect both responses by comparing the transcriptional changes that occur in a wild-type strain and in a hetR mutant strain that is unable to differentiate heterocysts. The construction of a co- expression network followed by a clustering analysis allowed us to define ten groups of genes according to their expression profiles (Chapter III, Figure S1).

Two of the clusters included genes directly regulated by NtcA (Mitschke et al., 2011b; Picossi et al., 2014) and involved in the general response to nitrogen deprivation, as glnA, gifA or nblA (see section 1.1.4.1). In addition, we identified two clusters containing elements (protein coding genes, sRNAs, etc) whose expression was HetR-dependent and, thus, could be specifically involved in heterocyst differentiation (Chapter III, Figure 1). Transcription of the elements in these two clusters showed different temporal profiles, corresponding to an early (E-DIF) or late (L-DIF) expression along the differentiation process. Many genes grouped in these two clusters had been previously identified in screenings of mutants unable to differentiate functional

219 General discussion

heterocysts. Our approach allowed the identification of additional elements that remained unnoticed either because it is not possible to mutate them, or because their functions are redundant and, consequently, they would not yield a distinct phenotype when mutated. However, based on their clustering with known genes related to heterocyst development, they are also likely involved in the heterocyst differentiation process.

For previously described genes, their inclusion in the early (E-DIF) or late (L-DIF) clusters was consistent with data in the literature. For instance, genes in the “hep island” were expressed earlier (E-DIF) than genes in the “hgl island” (L-DIF) (Chapter III, Figure S2), as previously reported (Ehira et al., 2003; Flaherty et al., 2011). Among the genes involved in the generation and maintenance of patterning of heterocysts, patS (Yoon and Golden, 1998), patC (Corrales-Guerrero et al., 2014), and patX (Elhai and Khudyakov, 2018) were expressed earlier than hetN (Callahan and Buikema, 2001) in our transcriptional profiles. These results are consistent with the different roles proposed for PatS and HetN in the de novo generation or maintenance of the pattern, respectively (Rivers et al., 2018). In addition, different temporal expression was also shown for genes related to oxygen protection. rbrA (Zhao et al., 2007), encoding a rubrerythrin, and alr3808 (Li et al., 2015), encoding a Dps protein, were expressed earlier than the cytochrome oxidases encoded in the cox2 and cox3 operons (Valladares et al., 2003) or the flavodiiron proteins encoded in the flv1B and flvB3 genes (Ermakova et al., 2014).

As previously mentioned, the microarrays used for the hybridization experiments leading to the construction of the co-expression network included probes for putative sRNAs and asRNA whose TSS had been identified in a previous dRNAseq experiment (Mitschke et al., 2011b). Interestingly, many sRNAs and asRNA were included in the clusters associated to heterocyst differentiation. The presence of many RNA regulators in the clusters associated to heterocyst differentiation may suggest an important role of these molecules in the regulation of this differentiation process. For instance, the heterocyst-specific, HetR-dependent asRNA As_glpX, antisense to the gene alr1041, included in the E-DIF cluster seems to contribute to the inhibition of the Calvin cycle in heterocysts (Olmedo-Verd et al., 2019). Other non-coding RNAs whose HetR-dependent expression was verified were the asRNA As_alr5059 (Chapter III, Figure 2) and the sRNA NsiR2 (Chapter III, Figure S3) both expressed early (E-DIF cluster), and the sRNA NsiR9 (Chapter III, Figure S3) that was expressed late in heterocyst development (L-DIF cluster).

We detected some DNA motifs associated to cell-specific expression. Transcripts in the E-DIF cluster showed a relatively homogeneous profile of expression that pointed to a limited number or regulating factors. We found an enrichment of the previously described DIF1 motif

220 General discussion

(Mitschke et al., 2011b; Muro-Pastor, 2014; Muro-Pastor et al., 2017) and a new motif, DIF2 (Chapter III, Figure 4), in the promoters of elements grouped in the E-DIF cluster (Chapter III, Figure S5). However, we could not detect any new DNA motif that appeared enriched in the promoter sequences of elements in L-DIF cluster, but only detected the previously described CnfR motif (Vernon et al., 2017). So, in this work we have detected new genes and non-coding transcripts that may have a role in the differentiation of heterocyst and whose cell-specific and temporal expression may be driven by newly identified DNA motifs (Chapter III, Figure 5).

4.4 NsiR1, a sRNA with multiple copies, regulates heterocyst differentiation

NsiR1 is a sRNA of about 60 nucleotides that is induced at an early stage during the differentiation of cells becoming heterocysts (Muro-Pastor, 2014). It is encoded in multiple tandem copies in the genomes of Nostoc sp. PCC 7120 and other heterocyst-forming cyanobacteria (Ionescu et al., 2010). Whereas the central copies (#3-9, called NsiR1.4) are identical, the rest of the copies show some sequence divergence. In addition, due to their genomic location, one copy (NsiR1.1) may act as a perfect antisense to the 5’UTR of hetF, encoding a protease involved in the regulation of HetR and playing a relevant role in the differentiation of heterocysts.

Our results showed that there is a significant co-occurrence of nsiR1 and hetF in the genomes of heterocyst and non-heterocyst forming filamentous strains (Chapter IV, Figure S1). The number of copies of nsiR1 is higher in heterocyst-forming cyanobacteria and those copies are located in the region upstream of hetF homologs (Chapter IV, Figure S2). This sinteny analysis showed that the function of NsiR1.1 as antisense to hetF may be conserved along heterocyst- forming cyanobacteria.

Data obtained in E. coli showed that NsiR1.1 may negatively regulate the expression of hetF (Chapter IV, Figure 1D-E). This result was supported by the use of translational fusions of hetF 5’UTR to gfpmut2 under the control of the rnpB promoter in Nostoc, showing a negative post-transcriptional regulation of hetF in heterocysts, that could be due to the negative effect of NsiR1.1 (Chapter IV, Figure 1C). In contrast to many other described sRNAs, the predicted interaction of NsiR1.1 with hetF 5’UTR was located far from the translation initiation region. In vitro footprinting assays showed a conformational change in the hetF 5’UTR in the presence of NsiR1.1 that provokes a protection in the translation initiation region of hetF (Chapter IV, Figure 2), which may explain the negative regulation exerted by NsiR1.1 on HetF translation.

221 General discussion

The same strategy used for the expression and depletion of Yfr1 (Chapter II) was used for NsiR1. We generated strains that overexpressed NsiR1.1, NsiR1.4 or a sequence antisense to NsiR1.4 that lead to depletion of the pool of endogenous NsiR1 (Chapter IV, Figure 3B). Because we were unable to detect intact hetF mRNA, and because HetF is a protease that degrades HetR (Risser and Callahan, 2008) we measured the accumulation of HetR as an indirect indication of HetF function. We detected a higher amount of HetR in the OE_NsiR1.1 strain (Chapter IV, Figure 3C-D) that was not due to a transcriptional induction of hetR (Chapter IV, Figure S4) and was consistent with a negative regulation of hetF expression by NsiR1.1.

The phenotypic characterization of Nostoc strains with altered levels of NsiR1 showed a functional divergence between NsiR1.1 and NsiR1.4. Although these two versions of NsiR1 only differ in several nucleotides, the strain that overexpressed NsiR1.4 differentiated heterocysts in media with nitrate (and this was not due to a lack of nitrate assimilation), whereas the strain that overexpressed NsiR1.1 did not differentiate heterocysts under this condition (Chapter IV, Figure 4). Therefore, NsiR1.4 seemed to favour heterocyst differentiation, which is coherent with the observation that the strain with less NsiR1 showed a delay in the differentiation process (Chapter IV, Figure 4D).

A prediction of targets indicated that NsiR1.4 may interact with alr3234 mRNA (Ionescu et al., 2010). This interaction was validated in E. coli and by in vitro footprinting assays (Chapter IV, Figure 5). In this case, the interaction site overlapped the alr3234 start codon, suggesting a negative regulation based on the blocking of translation initiation. This putative negative regulation was also validated in Nostoc using the same strategy carried out with hetF; a translational fusion of alr3234 5’UTR to gfpmut2 showed a negative post-transcriptional regulation of alr3234 in heterocysts, that could be due to the negative effect of NsiR1.4 (Chapter IV, Figure 6A). It has been described that in an alr3234 mutant the timing of commitment to heterocyst differentiation is shifted to earlier time points with respect to the wild type strain (Videau et al., 2016a). We performed commitment assays that showed that the OE_NsiR1.4 strain in fact had earlier commitment than the control strain (Chapter IV, Figure 6B), whereas the OE_as_NsiR1 strain had a slightly delayed commitment to heterocyst differentiation, in agreement with the proposed negative regulation of alr3234 by NsiR1.4.

Sibling sRNAs (molecules with similar sequences encoded in a given genome) have been described in different bacteria (Caswell et al., 2014) but most of these sibling sRNAs have redundant functions. In our case, although the different versions of NsiR1 may have redundant targets, we have validated a functional divergence between NsiR1.1 and NsiR1.4 that may allow

222 General discussion these two versions of NsiR1 to regulate different aspects of a global process. In this case, NsiR1.1 and NsiR1.4 would regulate heterocyst differentiation by increasing the local amount of HetR (NsiR1.1) and modifying timing of commitment (NsiR1.4). In addition, NsiR1 is the first described sRNA involved in heterocyst differentiation. So, our results point to an important role of RNA regulators in the regulation of heterocyst differentiation through a complex regulatory network based on the interplay between classical transcriptional regulation exerted by transcription factors and post-transcriptional regulation exerted by other mechanisms, including sRNA regulation (Chapter IV, Figure 7).

223

5. CONCLUSIONS

Conclusions

5. CONCLUSIONS

1. A computational approach based on dRNAseq data, a prediction of Rho-independent terminators and phylogenetic conservation identifies 327 possible sRNA genes in the genome of Nostoc sp. PCC 7120. 2. Some of the predicted sRNA genes are only conserved in heterocystous cyanobacterial genomes, suggesting a putative role of these sRNAs in the regulation of the specific metabolism or physiology of these cyanobacteria. 3. Yfr1 regulates the expression of genes encoding outer membrane proteins or enzymes related to peptidoglycan biosynthesis and remodeling in Nostoc sp. PCC 7120. 4. Strains with altered levels of Yfr1 have an altered outer membrane and an unusual distribution of septa and nascent peptidoglycan. 5. The overexpression of Yfr1 prevents diazotrophic growth probably due to disruption of the connections between vegetative cells and heterocysts. 6. The use of a co-expression network allows the dissection of the complex dynamic changes in the transcriptome of Nostoc sp. PCC 7120, clustering together genetic elements with similar expression profiles. 7. Two clusters of transcripts (E-DIF and L-DIF) with different temporal profiles were associated to heterocyst-differentiation. Some RNA regulators were found in these clusters and their cell-specific expression was validated. 8. Some DNA sequence motifs (DIF1, DIF2 and CnfR motifs) are enriched among the promoter sequences of genetic elements that are specifically expressed in heterocysts. 9. Two heterocyst-specific sibling RNAs have different regulatory targets. NsiR1.1 represses the expression of hetF whereas NsiR1.4 represses the expression of alr3234. 10. The cell-specific expression of NsiR1.1 and NsiR1.4 in heterocysts, plus the constitutive transcriptional expression of hetF and alr3234 in all cells, generates a complex regulatory pattern in which the expression of hetF and alr3234 would be downregulated only in heterocysts.

227

6. GENERAL REFERENCES

General references

6. GENERAL REFERENCES

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