BIOTECHNOLOGY OF MICROALGAE, BASED ON MOLECULAR BIOLOGY AND BIOCHEMISTRY OF EUKARYOTIC ALGAE AND CYANOBACTERIA

EDITED BY: Takashi Osanai, Youn-Il Park and Yuki Nakamura PUBLISHED IN: Frontiers in Microbiology

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ISSN 1664-8714 ISBN 978-2-88945-129-6 DOI 10.3389/978-2-88945-129-6

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# **BIOTECHNOLOGY OF MICROALGAE, BASED ON MOLECULAR BIOLOGY AND BIOCHEMISTRY OF EUKARYOTIC ALGAE AND CYANOBACTERIA**

Topic Editors: **Takashi Osanai,** Meiji University, Japan **Youn-Il Park,** Chungnam National University, South Korea **Yuki Nakamura,** Academia Sinica, Taiwan

**Citation:** Osanai, T., Park, Y-Il., Nakamura, Y., eds. (2017). Biotechnology of Microalgae, Based on Molecular Biology and Biochemistry of Eukaryotic Algae and Cyanobacteria. Lausanne: Frontiers Media. doi: 10.3389/978-2-88945-129-6

# Table of Contents

Hiroko Iijima, Tomokazu Shirai, Mami Okamoto, Akihiko Kondo, Masami Yokota Hirai and Takashi Osanai

Jong-Min Lim, Jayaraman Vikramathithan, Kwon Hwangbo, Joon-Woo Ahn, Youn-Il Park, Dong-Woog Choi and Won-Joong Jeong

# Editorial: Biotechnology of Microalgae, Based on Molecular Biology and Biochemistry of Eukaryotic Algae and Cyanobacteria

Takashi Osanai 1 \*, Youn-Il Park 2 \* and Yuki Nakamura3 \*

1 School of Agriculture, Meiji University, Kawasaki, Japan, 2 Department of Biological Sciences, Chungnam National University, Daejeon, South Korea, 3 Institute of Plant and Microbial Biology, Academia Sinica, Taipei, Taiwan

Keywords: Chlamydomonas, Chlorella, cyanobacteria, metabolic engineering, microalgae, photoreceptor, Synechocystis, triacylglycerol

**Editorial on the Research Topic**

#### Edited by:

William James Hickey, University of Wisconsin-Madison, USA

#### Reviewed by:

Regina-Michaela Wittich, Spanish High Council for Scientific Research - Estación Experimental del Zaidín, Spain

#### \*Correspondence:

Takashi Osanai<EMAIL_ADDRESS>Youn-Il Park<EMAIL_ADDRESS>Yuki Nakamura<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology

> Received: 18 November 2016 Accepted: 17 January 2017 Published: 01 February 2017

#### Citation:

Osanai T, Park Y-I and Nakamura Y (2017) Editorial: Biotechnology of Microalgae, Based on Molecular Biology and Biochemistry of Eukaryotic Algae and Cyanobacteria. Front. Microbiol. 8:118. doi: 10.3389/fmicb.2017.00118

### **Biotechnology of Microalgae, Based on Molecular Biology and Biochemistry of Eukaryotic Algae and Cyanobacteria**

Microalgae, including eukaryotic algae and cyanobacteria, comprise a diverse group of microscopic unicellular photosynthetic organisms that inhabit almost all ecological niches. Recently, energy and resource production using microalgae have received a great deal of attention (Banerjee et al., 2016). Several species of microalgae are commercially available. They include Arthrospira platensis, Chlorella species, and Euglena gracilis (Koller et al., 2014; Gouveia et al., 2016; Yamada et al., 2016). Microalgae produce high-value products such as pigments, lipids, bioplastics, carbohydrates, and amino acids (Minhas et al., 2016). However, production by microalgae is generally less efficient than current industrial production. To increase productivity, a basic understanding of photosynthesis, metabolism, and the cellular structure of microalgae using molecular genetics is indispensable. Here, we introduce current researches, which cover the basic and applied science of eukaryotic algae and cyanobacteria.

The study of the primary carbon metabolism of photosynthetic organisms has a long history, and basic scientists have devoted their time to deciphering the regulatory mechanisms of primary metabolism and the biochemical properties of metabolic enzymes. Basic scientists have studied the regulator mechanisms of anabolic carbon processes such as carbon fixation, gluconeogenesis, and glycogen/starch production. However, the regulatory mechanisms of the catabolic processes of microalgae have been less well studied. The downstream metabolites of sugar catabolism are the precursors of biofuels and bulk chemicals, and an understanding of metabolic sugar regulation would therefore lead to improvements in productive capacity and chemical variety (Oliver et al., 2016). Unicellular cyanobacteria, such as Synechocystis and Synechococcus species, and filamentous cyanobacteria, such as Nostoc species, are the preferred microbial cell factories because they are easy to genetically manipulate (Atsumi et al., 2009; Khanna and Lindblad, 2015). Recently, there has been an increase in the number of publications dealing with the metabolic engineering of cyanobacteria. Several transcriptional regulators that activate the gene expression of sugar catabolic enzymes in cyanobacteria have been identified in Synechocystis; for example, an RNA polymerase sigma factor (SigE) and a response regulator (Rre37) (Osanai et al., 2011, 2014). Overexpression of these genes causes increased production of polyhydroxybutyrates, which are bioplastic polyesters (Osanai et al., 2014). Therefore, the elucidation of the regulatory mechanism of primary carbon metabolism would be important.

An understanding of photosynthetic electron transport systems and light-to-chemical energy conversion is indispensable for biorefineries. Varying light conditions mean that photoautotrophs must sense light intensity, quality, and light/dark photoperiods to carry out optimal photosynthesis (Eberhard et al., 2008). Near-UV light, which causes damage to the photosynthetic apparatus, is effectively screened-out by light-blocking pigments such as anthocyanin and other near-UV absorbing pigments. The application of multiple interdisciplinary approaches has expanded our knowledge of the structure and function of traditionally well-characterized photoreceptors and their downstream signaling mechanisms. The last 10 years in particular have witnessed discoveries in the newly emerging field of plant photobiology, especially bilin photoreceptor phytochromes (Rockwell and Lagarias, 2010).

Various types of cyanobacterial phytochromes have been discovered using genomic information from Synechocystis, and heterocystous (Nostoc punctiforme) and non-heterocystous (Microcoleus IPPAS B353) cyanobacteria; they are distinguished by their domain structures and by the number of cysteine (Cys) amino acid residues they contain, which play a role in the attachment of tetrapyrrole chromophores. Cyanobacteria are able to sense near-UV and the entire visible light range using various bilin photoreceptors. Their biological functions are implicated in positive and negative phototaxis, state transition, and salt acclimation, and to a lesser extent in other photobiological responses such as carotenoid accumulation and filament stacking (Wiltbank and Kehoe, 2016). Moreover, photosynthesis is regulated by photoreceptors such as phototropin from the green algae Chlamydomonas reinhardtii (Petroutsos et al., 2016) and cyanobacterial phytochromes Cph2 from Synechocystis (Schwarzkopf et al., 2014); the discovery of these photoreceptors has paved the way for the study of the coevolution of lightsensing and signaling pathways. Some eukaryotic algae contain phytochromes that are responsive to the visible spectrum, rather than to canonical red/far red light (Rockwell et al., 2014), which is a fascinating aspect of phytochrome evolution. Basically, cyanobacterial bilin photoreceptors transduce signals to the immediate downstream component via phosphorylation. These phosphorylated components often act as transcriptional factors, activating the downstream cascade related to transition state and phototaxis. Using these two-component systems, optogenetic approaches can be applied to selectively regulate gene expression for nutrient uptake and augment particular metabolic pathways in the field of biotechnology (Narikawa et al., 2015).

The study of the regulatory mechanisms that maintain cellular structure is also important for both basic and applied science.

### REFERENCES

Lipids are the major constituents of cellular membranes (Li-Beisson et al., 2016). More specifically, microalgae contain thylakoid membranes, which are the sites of photosynthetic reactions that require the absorption of light energy to split water, and produce oxygen, ATP, and reductants. Moreover, lipids can be converted into biofuels. Microalgae are known for their high capacity for lipid production. Triacylglycerols and other neutral lipids are the major form of storage lipids; they are often termed "oils," and represent an inert end-product of photosynthetic carbon assimilation. Compared with the higher plants in which storage lipid accumulation only occurs in limited types of tissues/cells, unicellular microalgae have a high degree of flexibility that alters cellular lipid metabolic flux (i.e., lipid biosynthesis for photosynthetic membrane constructions such as in plant leaves, or lipid accumulation for storage in plant seeds) during growth. For instance, nitrogen starvation alters the growth phase of algae and induces oil accumulation at the expense of cellular membrane lipids in many algal species. Thus, flexible lipid metabolic flux in unicellular systems, overall high lipid productivity, photosynthetic capacity that allows carbon fixation, and rapidly growing gene manipulation technologies, have promoted the use of microalgae as an emerging synthetic platform in metabolic engineering. A basic understanding of complex algal lipid metabolism will guide us to the most feasible and effective strategies for creating biofuels.

Therefore, microalgae have great potential for biofuel and biomaterial production. However, this translational research requires knowledge of molecular biology and biochemistry. Therefore, we believe that these researches contribute to crossing the "valley of death" (the gap between basic research and commercialization) in the future.

### AUTHOR CONTRIBUTIONS

All the authors of this manuscript confirmed their contribution and approved it for publication.

### ACKNOWLEDGMENTS

This work was supported by the Ministry of Education, Culture, Sports, Science, and Technology, Japan; by a grant to TO from ALCA (Project name "Production of cyanobacterial succinate by the genetic engineering of transcriptional regulators and circadian clocks") from the Japan Science and Technology Agency; and by the JSPS KAKENHI Grant-in-Aid for Scientific Research on Innovative Areas Grant Number 16H06559.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Osanai, Park and Nakamura. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# A Screening Method for the Isolation of Polyhydroxyalkanoate-Producing Purple Non-sulfur Photosynthetic Bacteria from Natural Seawater

Mieko Higuchi-Takeuchi, Kumiko Morisaki and Keiji Numata\*

Enzyme Research Team, Biomass Engineering Research Division, RIKEN Center for Sustainable Resource Science, Wako, Japan

Polyhydroxyalkanoates (PHAs) are a family of biopolyesters accumulated by a variety of microorganisms as carbon and energy storage under starvation conditions. We focused on marine purple non-sulfur photosynthetic bacteria as host microorganisms for PHA production and developed a method for their isolation from natural seawater. To identify novel PHA-producing marine purple non-sulfur photosynthetic bacteria, natural seawaters were cultured in nutrient-rich medium for purple non-sulfur photosynthetic bacteria, and twelve pink- or red-pigmented colonies were picked up. Gas chromatography mass spectrometry analysis revealed that four isolates synthesized PHA at levels ranging from 0.5 to 24.4 wt% of cell dry weight. The 16S ribosomal RNA sequence analysis revealed that one isolate (HM2) showed 100% identity to marine purple non-sulfur photosynthetic bacteria. In conclusion, we have demonstrated in this study that PHA-producing marine purple non-sulfur photosynthetic bacteria can be isolated from natural seawater under nutrient-rich conditions.

Keywords: polyhydroxyalkanoates, marine purple non-sulfur photosynthetic bacteria, seawater, nutrient-rich conditions, 16S rRNA

## INTRODUCTION

Polyhydroxyalkanoates (PHAs) are biopolyesters that many microorganisms accumulate as intracellular reservoirs of carbon and energy. PHAs have attracted increasing attention due to their biomass-based origin and biodegradability (Numata et al., 2009). However, PHA production requires a costly carbon source, such as sugars or plant oils. The use of photosynthetic organisms to produce materials is clean and eco-friendly because the energy for growth is derived from the sun, and the carbon is derived from carbon dioxide in the air. Although efforts have been made to utilize photosynthetic organisms such as cyanobacteria and higher plants for PHA production, high PHA production has not been achieved to date (Osanai et al., 2013, 2014). Anoxygenic photosynthetic bacteria are known to produce more PHA (Liebergesell et al., 1991) than cyanobacteria and higher plants. However, PHA production by photosynthetic bacteria has been studied in a small number of freshwater purple non-sulfur photosynthetic bacteria strains, such as Rhodospirillum rubrum (Brandl et al., 1989; Clemente et al., 2000), Rhodopseudomonas sphaeroides (Lorrungruang et al., 2006) and Rhodobacter capsulatus (Kranz et al., 1997). Purple non-sulfur photosynthetic bacteria have several advantages over other photosynthetic bacteria. One advantage is that purple non-sulfur photosynthetic bacteria can grow either aerobically in the dark or anaerobically in the

#### Edited by:

Yuki Nakamura, Institute of Plant and Microbial Biology (Academia Sinica), Taiwan

#### Reviewed by:

Kesaven Bhubalan, Universiti Malaysia Terengganu, Malaysia Vidya De Gannes, The University of the West Indies– St. Augustine Campus, Trinidad and Tobago

> \*Correspondence: Keiji Numata<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology

Received: 27 July 2016 Accepted: 09 September 2016 Published: 21 September 2016

#### Citation:

Higuchi -Takeuchi M, Morisaki K and Numata K (2016) A Screening Method for the Isolation of Polyhydroxyalkanoate-Producing Purple Non-sulfur Photosynthetic Bacteria from Natural Seawater. Front. Microbiol. 7:1509. doi: 10.3389/fmicb.2016.01509

light. In addition, purple non-sulfur photosynthetic bacteria can utilize various electron acceptors, i.e., they are facultative photosynthetic bacteria (Mcewan, 1994; Basak and Das, 2007). To take advantage of these properties, purple non-sulfur photosynthetic bacteria have been tested for use in a variety of applications, including not only PHA production but also the purification of industrial wastewater (Kim et al., 2004; Wu et al., 2012) and hydrogen production (Basak and Das, 2007).

Marine microorganisms are important bioresources and expected to produce new value-added compounds, including PHA (Numata and Doi, 2012; Numata et al., 2013; Numata and Morisaki, 2015). Cultivation under marine conditions offers several advantages for the industrial production of PHA. For example, high concentrations of salts inhibit the growth of saltsensitive bacteria species. In addition, filtered sterilized seawater can be used as a culture medium. However, PHA production under marine conditions has been reported using certain types of marine bacteria (Lopez et al., 2009; Shrivastav et al., 2010; Numata and Doi, 2012). Although PHA production by marine purple non-sulfur photosynthetic bacteria has been reported by a few groups (Chowdhury et al., 1996; Xiao and Jiao, 2011), the details of the PHA synthesis were not studied thoroughly. The small number of studies on PHA synthesis by marine purple nonsulfur photosynthetic bacteria, even though they are important host bacteria to produce PHA, is because no isolation method of PHA-producing marine purple non-sulfur photosynthetic bacteria has been established until now.

Many screening methods have been developed to detect microorganisms that accumulate PHAs. The methods most widely used for detecting PHAs are staining techniques using Nile red (Spiekermann et al., 1999), Nile blue A (Ostle and Holt, 1982) and Sudan Black (Steinbuchel et al., 1987). Using these staining techniques, a variety of PHA-producing bacteria and mutants have been isolated. However, this method is unable to discriminate PHA and lipids. Additionally, it is necessary to provide nutrient limitation conditions and suitable carbon sources to the bacterial cells to induce PHA production. Purple photosynthetic bacteria contain bacteriochlorophyll a or b and various types of carotenoids. Staining methods are not suitable for purple non-sulfur photosynthetic bacteria because these pigments interfere with staining and detection. The other method for identifying PHA-producing bacteria is the polymerase chain reaction (PCR) amplification of PHA synthesis genes using degenerate primers (Sheu et al., 2000; Shamala et al., 2003). This technique is a rapid and accurate detection system for screening large numbers of environmental isolates. However, this technique leads to detection errors because of the nonspecific PCR amplification and lack of PCR products due to degenerate primer sequences. Furthermore, this method cannot detect PHA itself, and hence PHA and its induction conditions must be determined after the discovery of PHA biosynthesis genes. Therefore, it is necessary to develop a screening method for the isolation of PHA-producing marine purple non-sulfur photosynthetic bacteria.

In a previous study, we evaluated the potential use of marine purple photosynthetic bacteria for PHA production (Higuchi-Takeuchi et al., 2016). Our study demonstrated that marine purple photosynthetic bacteria were good host microorganisms for industrial PHA production using marine resources. The aim of our study is to develop an approach to isolate PHAproducing purple photosynthetic bacteria from natural marine environments. We found that purple sulfur photosynthetic bacteria did not accumulate PHA under nutrient-rich conditions, whereas some species of purple non-sulfur photosynthetic bacteria did accumulate PHA without nutrient limitation, in contrast to the well-known PHA-producing soil bacteria (Higuchi-Takeuchi et al., 2016). Purple photosynthetic bacterial cultures can be purple, red, brown or orange because of the various types of carotenoids and bacteriochlorophyll. Based on these observations, in this study, we isolated pigmented bacteria under nutrient-rich conditions as PHA-producing purple nonsulfur photosynthetic bacteria from natural seawaters.

## MATERIALS AND METHODS

### Culture Conditions and Seawater Sampling

Purple non-sulfur bacteria were grown in culture medium (JCM medium number 520)1 . The composition was modified based on the medium used for isolation of purple non-sulfur bacteria (Biebl and Pfennig, 1981). Medium was composed of the following components per liter: KH2PO4 (0.5 g); CaCl2·2H2O (0.25 g); MgSO4 ·7H2O (3.0 g); NH4Cl (0.68 g); NaCl (20 g); sodium malate (3.0 g); sodium pyruvate (3.0 g); yeast extract (0.4 g); ferric citrate (5 mg); vitamin B12 (2 mg); ZnCl2·5H2O (70 µg); MnCl2·4H2O (100 µg); H3BO3 (60 µg); CoCl2·6H2O (200 µg); CuCl2·2H2O (20 µg); NiCl2·6H2O (20 µg) and Na2MoO4·H2O (40 µg). The pH was adjusted to 6.8. Yeast extract (Culture media grade) and ferric citrate (BioReagent grade) were purchased from Becton Dickinson (Franklin Lakes, NJ, USA) and Sigma-Aldrich (St. Louis, MO, USA), respectively. The rest chemicals (JIS Special Grade) were purchased from Wako Chemicals (Osaka, Japan).

For screening, aliquots of 100 µL of each seawater sample from all sampling points were spread in agar plates. Seawaters were cultured multiple times until pigmented-colony was obtained. Maximum number of replicates was six. Six replicates were carried out using seawaters from Naha, Tokyo-Bay and Sakata. The number of replicates was five in Omura-Bay sample, six in Tokyo-Bay, four in Aioi-Bay, two in Takamatsu and two in Yokohama. Seawaters were cultured aerobically under continuous far-red LED light conditions (730 nm, 8 Wm−2 ) at 30◦C. Addition of larger amount of seawater and concentration of seawater using filter paper might be effective for obtaining the large number of bacterial colonies. The isolated colonies were streaked onto other plates for purification. Streak plating should be done until the pure colonies from single bacteria are isolated. Transferring of small volume of cell culture (1.5 mL) into large scale culture (50 mL) is recommended for better bacterial growth. The isolates were transferred to 1.5 mL capped plastic tubes filled with liquid medium and cultured for 7–10 days under continuous

1http://www.jcm.riken.jp/cgi-bin/jcm/jcm\_grmd?GRMD=520

far-red LED light conditions at 30◦C and then transferred in 50 mL screw capped plastic tubes filled with liquid medium and cultured without stirring for 10 days. To transfer into liquid culture, transferring a small amount of bacteria cells (not a single colony) using inoculating loops under aseptic conditions is preferred for good bacterial cell growth. Bacterial cells were stored as 10% glycerol (JIS Special Grade, Wako Chemicals, Osaka, Japan) stocks at −80◦C for further experiments.

About 50 mL of natural seawaters were collected from Naha (Okinawa), Omura-Bay (Nagasaki), Tokyo-Bay (Tokyo), Aioi-Bay (Hyogo), Sakata (Yamagata), Takamatsu (Kagawa) and Yokohama (Kanagawa) in Japan. Sampling points were randomly selected. One seawater sample was collected from the sea surface in each point. Sampled seawaters were stored at 4◦C in the dark until use.

### Analysis of PHA Content and Composition

The PHA content characterization method was modified slightly from a previous study (Chuah et al., 2013). Approximately 0.5–2 mg lyophilized cells were incubated in 1 mL of 100% ethanol at 70◦C for 1 h to remove pigments. The cells were then subjected to ethanolysis in the presence of 250 µL chloroform (JIS Special Grade, Wako Chemicals, Osaka, Japan), 100 µL hydrochloric acid (JIS Special Grade, Wako Chemicals, Osaka, Japan) and 850 µL ethanol (JIS Special Grade, Wako Chemicals, Osaka, Japan) at 100◦C for 4 h. After cooling, 1 mL of phosphate buffer (pH 8.1) was added to the reaction mixture and then neutralized with 0.65 N NaOH. After centrifugation at 1,500 rpm for 5 min (CF16RN, Hitachi-Koki, Tokyo, Japan), the lower chloroform layer was filtered through anhydrous sodium sulfate (JIS Special Grade, Wako Chemicals, Osaka, Japan) and incubated with molecular sieves 4A (Nacalai tesque, Kyoto, Japan) for 30 min. The PHA content and composition were determined using a gas chromatography–mass spectrometry (GC-MS) apparatus (GCMS-QP2010 Ultra, Shimadzu, Tokyo, Japan) equipped with a 30 mm × 0.25 mm DB-1 capillary gas chromatography column (Agilent Technologies, Santa Clara, CA, USA). For analysis, 1 µL of sample solution was injected with helium as a carrier gas (3.30 mL min−1 ). The following temperature program was used to separate ethyl esters: 45◦C for 1 min, temperature ramp of 7◦C per min to 117◦C. The interface and ion source temperatures were 250◦C and 230◦C, respectively. The 3HB content was determined using a calibration curve.

### Extraction of PHAs and 1H NMR analysis

PHA extraction and purification was carried out according to the method reported in (Brandl et al., 1989). PHAs were extracted from 428 mg lyophilized cells using about 100 mL of chloroform (JIS Special Grade, Wako Chemicals, Osaka, Japan). The chloroform extracts were filtered and concentrated using a rotary vacuum evaporator (NA-1, AS ONE, Osaka, Japan), and the chloroform-extracted PHAs were purified by precipitation with hexane (JIS Special Grade, Wako Chemicals, Osaka, Japan) of more than 10 times the volume of the solvent. The precipitate was filtered and then air-dried without vacuum as a convenient way overnight at room temperature. The polymer solution was concentrated again using a rotary vacuum evaporator and purified by precipitation with cold methanol (JIS Special Grade, Wako Chemicals, Osaka, Japan) of more than 10 times the volume of the solvent. The purified PHA precipitate was air-dried overnight at room temperature.

The purified PHAs were analyzed by proton nuclear magnetic resonance (1H NMR; JNM-Excalibur 270; JEOL, Ltd., Tokyo, Japan) to determine their chemical structures and compositions. The measuring frequency was 499.87 MHz. The sample for NMR analysis was dissolved at a concentration of 4 mg/mL in CDCl3 with 0.05% (v/v) tetramethylsilane (TMS; Wako Pure Chemical Industries Ltd., Osaka, Japan).

### Determination of 16S Ribosomal RNA Sequences and Phylogenetic Analysis

DNA was extracted using the DNeasy Blood & Tissue Kit (Qiagen, Hilden, Germany). The 16S ribosomal RNA sequences were determined using the Bacterial 16S rDNA PCR kit according to the manufacturer's protocol (Takara Bio, Shiga, Japan).

The phylogenetic tree was prepared based on the 16S ribosomal RNA sequences of the 13 selected purple photosynthetic bacteria and constructed using Phylogeny2 (Castresana, 2000; Edgar, 2004; Anisimova and Gascuel, 2006; Dereeper et al., 2008; Guindon et al., 2010). NJplot was used to display the phylogenetic tree (Perriere and Gouy, 1996).

### RESULTS AND DISCUSSION

The screening strategy for PHA-producing purple non-sulfur photosynthetic bacteria is shown in **Figure 1**. In the case of well-known PHA-producing bacteria, PHA production is induced under nutrient(s)-free conditions such as nitrogen, phosphorus or sulfur. We previously found that purple nonsulfur photosynthetic bacteria could accumulate PHA without nutrient deficiency (Higuchi-Takeuchi et al., 2016). Nutrient deficiency conditions are not appropriate for the isolation of PHA-producing purple non-sulfur photosynthetic bacteria from natural environments because such conditions result in poor bacterial growth. Therefore, seawaters were cultured in nutrient-rich agar plates for the growth of marine purple non-sulfur photosynthetic bacteria. To select PHA-producing purple non-sulfur photosynthetic bacteria by color, the plates

n.d., not detected.

FIGURE 2 | 1H-NMR spectra of purified PHA. 1H-NMR analysis was conducted using purified PHA from HM2 cells.

were cultivated under continuous far-red light conditions. After extended cultivation, a red- or pink-pigmented colony typical of purple photosynthetic bacteria was picked up. The selected colony was cultured in liquid medium, and the harvested cells were subjected to GC-MS analysis to measure their PHA content. Lastly, the 16S rRNA sequences were determined to identify the bacterial species.

To obtain purple non-sulfur photosynthetic bacteria colonies, each seawater sample from all sampling points was spread in nutrient-rich agar plates for the growth of purple non-sulfur

2http://phylogeny.lirmm.fr/phylo\_cgi/index.cgi

photosynthetic bacteria and cultured aerobically in the light. Most bacterial colonies appeared white, cream or yellow in color after 1 day (Supplementary Figure 1C). Some bacteria produced pink or red colonies after 4–7 days (Supplementary Figures 1A,B). Extended incubation times may permit the appearance of more pigmented-colonies although they result in growth of bacteria other than purple non-sulfur photosynthetic bacteria. A total of twelve red or pink colonies were obtained from four seawater samples (**Table 1**). HM1 and HM2 were obtained from the seawater of Omura-Bay; HM3 and HM4 were obtained from the seawater of Takamatsu; HM5 was obtained from the seawater of Aioi-Bay; and seven isolates (HM5 to HM12) were isolated from the seawater of Yokohama. Pigmented colonies were streaked onto other plates for further isolation (Supplementary Figures 1D–G). The twelve isolates were then transferred to liquid medium and cultured in the light (Supplementary Figures 1H–K). Among the twelve isolates, HM2 exhibited a bright red color in liquid culture (Supplementay Figure 1I). The cell cultures of the other eleven isolates were red in color after centrifugation (right side of Supplementary Figures 1H,J,K), although the liquid culture appeared to exhibit a very faint pink color or no color production (left side of Supplementary Figures 1H,J,K). Based on the results of the liquid cell cultures, HM2 was the only positive strain, namely, purple photosynthetic bacteria.

Twelve isolates were cultured in growth medium and then measured the cell dry weight (CDW) and PHA content. CDW varied among the isolates, ranging from 46 to 184 mg/L culture (**Table 1**). The PHA content of the twelve isolates was measured by GC-MS. PHA was not detected in eight isolates. Four isolates (HM1, HM2, HM3 and HM6) synthesized PHA at levels ranging from 0.5 to 24.4 wt% of CDW (**Table 1**). HM2 showed the highest PHA accumulation (24.4 wt%). To verify these results, the PHA content and CDW of HM2 were measured using triplicate cultures. The CDW of HM2 was 800 ± 58 mg/L culture, and the PHA content was 24.2 ± 1.8 wt%. GC-MS analysis also revealed that four isolates synthesized a homopolymer of 3-hydroxybutyrate (3HB), namely, poly(3-hydroxybutyrate) (PHB). The chemical structure of the synthesized PHAs was determined by 1H NMR with chloroform extracts from HM2 isolate. Methyl, methylene, and methine protons of 3HB units were detected by 1H NMR (**Figure 2**). Thus, the 1H NMR spectra confirmed the chemical structure of PHB.

The 16S ribosomal RNA sequences of four isolates were determined and compared with the databank contents using Nucleotide BLAST. Two isolates (HM3 and HM6) showed more than 99% sequence identity to Shewanella basaltis, and one isolate (HM1) exhibited high similarity to Marinobacter guineae (**Table 2**). S. basaltis (Chang et al., 2008) and M. guineae (Montes et al., 2008) are marine bacteria but not purple nonsulfur photosynthetic bacteria. The isolate HM2, which was positive by color selection in liquid culture and accumulated the largest amount of PHA, showed 100% identity to the purple non-sulfur photosynthetic bacteria Afifella marina (Imhoff, 1983)

FIGURE 3 | Phylogenetic tree of 16S ribosomal RNA gene from HM2 and purple non-sulfur photosynthetic bacteria. The scale bar shows the number of nucleotide substitutions per site. GenBank accession numbers are given in parentheses. The bootstrap values are shown at branch nodes. Allochromatium vinosum was used as an outgroup.

and Rhodopseudomonas julia (Kompantseva, 1989) isolated from saline environments, indicating that bright pigmented-color in liquid culture is one of important points for isolation of purple non-sulfur bacteria. A phylogenetic tree was constructed based on the 16S rRNA sequences from HM2 and purple nonsulfur photosynthetic bacteria of the alphaproteobacteria that have been studied by whole-genome analysis (**Figure 3**). HM2 was positioned close to Blastochloris viridis in the phylogenetic tree. Further characterization of the isolates will be needed to determine the species.

The isolation method we described here identified the strain HM2, which showed the highest PHA production among twelve isolates, and the 16S rRNA sequences of this strain were shown to have high similarity to marine purple non-sulfur photosynthetic bacteria. Based on the data on PHA accumulation and CDW, the PHA content of HM2 was calculated to be 199 ± 16 mg/L culture. Brandl et al. (1989) reported production levels of 500 mg/L PHA in R. sphaeroides and 390 mg/L PHA in R. rubrum. HM2 showed similar levels of PHA production to these strains. Seawater sampling methods were not optimized in this study. Sampling points were randomly selected and only one seawater sample was collected from each sampling point. Further investigations for sampling methods are required to obtain a lot of marine purple non-sulfur photosynthetic bacteria from natural seawaters. Further optimization of factors such as the PHA induction conditions and screening of PHA-producing marine purple non-sulfur photosynthetic bacteria will allow the development of PHA production by photosynthetic organisms.

### CONCLUSION

One isolate identified in this study accumulated 24.4 wt% PHA, and 16S rRNA gene sequence analysis revealed that this strain showed high similarity to marine purple nonsulfur photosynthetic bacteria. Thus, we successfully developed

### REFERENCES

a screening method to isolate PHA-producing purple nonsulfur photosynthetic bacteria under nutrient-rich and farred light conditions from the natural environment. With this isolation method, PHA production by marine purple non-sulfur photosynthetic bacteria will be studied widely, leading to green and eco-friendly PHA production from carbon dioxide and marine resources.

### AUTHOR CONTRIBUTIONS

MH-T and KN conceived and designed the study. KM performed the experiments. MH-T analyzed the data, and MH-T and KN interpreted the data. MH-T drafted the manuscript, and KN and MK approved the manuscript.

### FUNDING

This work was supported by Grants-in-Aid for Scientific Research (MH-T), the Impulsing Paradigm Change through Disruptive Technologies Program (ImPACT) (KN) and the RIKEN Biomass Engineering Program (KN).

### ACKNOWLEDGMENT

We thank Mr. Ryota Sato and Miss. Nao Ifuku for the seawater sampling.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2016.01509

SigE during nitrogen starvation in Synechocystis sp. PCC 6803. DNA Res. 20, 525–535. doi: 10.1093/dnares/dst028

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Higuchi-Takeuchi, Morisaki and Numata. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Identification of Sporopollenin as the Outer Layer of Cell Wall in Microalga *Chlorella protothecoides*

Xi He, Junbiao Dai\* and Qingyu Wu\*

*MOE Key Laboratory of Bioinformatics, Center for Synthetic and Systems Biology, School of Life Sciences, Tsinghua University, Beijing, China*

*Chlorella protothecoides* has been put forth as a promising candidate for commercial biodiesel production. However, the cost of biodiesel remains much higher than diesel from fossil fuel sources, partially due to the high costs of oil extraction from algae. Here, we identified the presence of a sporopollenin layer outside the polysaccharide cell wall; this was evaluated using transmission electron microscopy, 2-aminoethanol treatment, acetolysis, and Fourier Transform Infrared Spectroscopy. We also performed bioinformatics analysis of the genes of the *C. protothecoides* genome that are likely involved in sporopollenin synthesis, secretion, and translocation, and evaluated the expression of these genes via real-time PCR. We also found that that removal of this sporopollenin layer greatly improved the efficiency of oil extraction.

#### *Edited by:*

*Youn-Il Park, Chungnam National University, South Korea*

#### *Reviewed by:*

*Arumugam Muthu, Council of Scientific and Industrial Research, India Won-Joong Jeong, Korea Institute of Bioscience and Biotechnology, South Korea*

#### *\*Correspondence:*

*Junbiao Dai<EMAIL_ADDRESS>Qingyu Wu<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology*

> *Received: 20 March 2016 Accepted: 22 June 2016 Published: 30 June 2016*

#### *Citation:*

*He X, Dai J and Wu Q (2016) Identification of Sporopollenin as the Outer Layer of Cell Wall in Microalga Chlorella protothecoides. Front. Microbiol. 7:1047. doi: 10.3389/fmicb.2016.01047* Keywords: biodiesel, cell wall, *Chlorophyta*, oil-extraction, sporopollenin

## INTRODUCTION

Global fossil fuel consumption has been increasing due to both population and economic growth (Lewis and Nocera, 2006), resulting in an energy shortage and a series of environmental problems such as global warming and effluent gas emissions (Chirinos et al., 2006). In response to these challenges, vigorous research programs are underway that to develop alternative biofuels, which are considered necessary for environmental and economic sustainability (Turner, 1999). Unfortunately, to date, biodiesel fuels obtained from plants and animals can't satisfy the existing demand for transport fuels. Microalgae appears to be a particularly promising candidate for use in biodiesel production because of its rapid growth rate, high oil production capacity, and lack of competition for agricultural land (Chisti, 2007). However, the price of algal biofuels remains much higher than that of conventional fossil fuel, due to the high production cost of both biomass and oil extraction (Demirbas and Fatih Demirbas, 2011). Chlorella protothecoides, a member of the Chlorophyta, can grow autotrophically when light is available. However, when C. protothecoides is grown in heterotrophic conditions (limited nitrogen and abundant glucose), it is able to reach very high cell densities (51.2 g L−1 ) and accumulate large amounts of lipids within cells (55.2%; Xu et al., 2006). It is considered to be a good model for research addressing commercial biofuel production. Transition from autotrophic condition to heterotrophic condition can reduce economic costs in biomass production. Nevertheless, oil extraction from C. protothecoides continues to be a significant challenge; it is highly energy consuming and because the cells are hard to break without harsh conditions such as the use of a bead beater (Xiao et al., 2015). Presumably, this resistance to cell lysis may result from a special structure and/or from the composition of the cell wall. Such speculations motivated us to analyze the composition of the cell walls of C. protothecoides.

Microalgal cell walls are complex and poorly understood. The chlorella intraspecies variation in cell walls can be dramatic and thus it is difficult to predict which of the compounds will be present in any one strain (Gerken et al., 2013). While some chlorella have only a single layer, others have two layers with the microfibrillar layer proximal to the cytoplasmic membrane and a thin mono or trilaminar outer layer (Yamada and Sakaguchi, 1982). Previous studies about the cell walls of different algae have indicated that only a limited number of chlorella species and other green algae are capable of synthesizing recalcitrant cell walls for protection from chemical or bacterial degradation. For example, some taxa can produce sporopollenin (Geisert et al., 1987; Ueno, 2009), a major component in the cell walls of spores; it has been reported in several genera of green algae, including Chlorella, Scenedesmus, Pediastrum, Chara, Prototheca, and Coelastrum (Burczyk and Czygan, 1983; Komaristaya and Gorbulin, 2006). In addition, Pore et al. found that acid and alkaline hydrolysis of C. protothecoides destroyed the cells, but could not destroy the cell wall components (Pore, 1984). Furthermore, the cell wall components were found to be resistant to acetolysis, which lead them to conclude the presence of sporopollenin. However, Lu et al. reported that they could generate protoplasts of C. protothecoides successfully using cellulase and snailase, which suggests the absence of sporopollenin (Lu et al., 2012). Therefore, it is arguable whether sporopollenin is present in C. protothecoides. Due to the recalcitrant nature of sporopollenin and sporopollenin-like materials, little is known about their definite chemical structure (Brooks and Shaw, 1978; Delwiche et al., 1989). Some studies have described unique structural features (Atkinson et al., 1972), and the conserved biogenesis pathway of sporopollenin has been predicted in some detail (Ríos et al., 2013).

In this paper, we report the discovery that the cell walls of C. protothecoides are resistant to the cell wall degradation enzymes. This suggests the presence of a protective layer that presumably prevents enzymes from accessing the wall components. We performed transmission electron microscopy (TEM), 2-aminoethanol treatment, acetolysis, and Fourier Transform Infrared Spectroscopy and provide evidence that this presumed extra layer exists and is composed of sporopollenin. Subsequently, we performed bioinformatics analysis of the sequenced C. protothecoides genome to identify genes that are likely involved in sporopollenin biogenesis and analyzed the expression of these genes with real-time PCR methods. In addition, we used a microfluidic device and monitored the propagation of single algal cells in detail. We found that these cells employ the typical Chlorella reproduction pattern and that their cell walls contain sporopollenin throughout the entire life cycle. We conclude that this sporopollenin is likely the primary obstacle to efficient oil extraction in this important model algal species.

### MATERIALS AND METHODS

### Strains and Culture Conditions

C. protothecoides sp. 0710 was cultured as described previously (Yan et al., 2011). Briefly, the autotrophic algae was grown at 28◦C with continuous illumination at 40 µmol·m−2 ·s −2 . The heterotrophic algae was grown in basal medium supplemented with 30 g L−1 glucose and 2.5 g L−1 yeast extract. Cells were incubated at 28◦C in flasks with shaking at 220 rpm.

### Enzymatic Treatment of Cells

Enzymatic treatment was performed as to previous study (Lu et al., 2012), with slight modifications. Log-phase Chlorella cells were harvested by centrifugation at 3000 rpm for 5 min, then the cell pellet was suspended in 25 mM Tris buffer (pH 6.0) containing the cell wall degrading enzymes and 0.6 M Dmannitol. Besides using the same combination of cellulase and snailase as described, a list of commercially available enzymes including cellulase (Sigma Cat. No. C1184 and Newprobe R-10), snailase (Newprobe), cellulysin (Calbiochem Cat. No. 219466), hemicellulase (Sigma Cat. No. H2125), pectinase (Sigma Cat. No. P2611), pectolyase (Sigma Cat. No. P3026), lysozyme (Sigma Cat. No. L6876), and zymolase (Zymoreseach Cat. No. E1005), were applied, either individually or in combination, so as to obtain the optimal digestion condition. Each treatment was kept at 30◦C for 16 h. The cells were then harvested for further analysis.

### Fluorescence Microscopy

Cells, both before and after disruption, were incubated in the enzyme solution containing 2% cellulase and 1% snailase in 0.6 M sorbitol and 0.6 M mannitol at 30◦C for 16 h. These cells were placed onto a clean glass slide and one drop of Calcofluor-white stain (Sigma Cat. No. P3543) and one drop of 10% potassium hydroxide were added sequentially to the slide. Following incubated for 1 min, cell wall fluorescence was examined under a confocal microscope (SP5, Leica).

### Cell Wall Extraction

Cells in the logarithmic growth phase were harvested by centrifugation at 3000 rpm for 5 min. The pellet was washed three times and resuspended in deionized water. Cell wall fragments were isolated as described previously (Hills, 1973; Matias and Beveridge, 2005). The cells were then transferred into a new 2 ml microfuge tube to which 0.3 g acid-washed glass beads (Sigma Cat. No. 18406) were added and processed for 30 times, for 30 s each time, in a Mini-Beadbeater (BioSpec Cat. No. 3110BXEUR) at 5000 rpm. An equal volume of 10% SDS was added, and the samples were boiled for 5 min, followed by centrifugation at 10,000 rpm for 5 min. The supernatant was then removed and the cell wall fragments which form the white fraction on the top of pellets, were isolated and washed three times in deionized water to remove excess SDS before use in subsequent experiments.

### Transmission Electron Microscopy

Transmission electron microscopic analysis was performed at the microscope facility in School of Life Sciences of Tsinghua University using a Hitachi H7650B transmission electron microscope following previously-described protocols (Burczyk and Hesse, 1981). Related measurements were taken using the metrics tool in illustrator.

### Acetolysis Assay

Acetolysis was carried out for 5 min at 95–100◦C in a mixture of acetic anhydride and concentrated sulfuric acid at a ratio of 9:1 (V/V). Cells were transferred to glacial acetic acid both before and after acetolysis to protect the acetic anhydride from breakdown.

### Fourier Transform Infrared Spectroscopy

For infrared absorption spectroscopic analysis, the cell wall fragments were first treated with the enzyme mixture as described in the previously-detailed sample preparation protocol to remove the polysaccharide layer of the cell wall. The samples were then processed by washing at 50◦C for 5 min followed by treatment with a chloroform-methanol mixture at a ratio of 2:1 (V/V) at 50◦C for 5 min. The process was repeated twice. Then the residues were washed sequentially in glacial acetic acid; 0.1 M sodium acetate; 1.0 M sodium hydroxide containing 0.01% tritonX-100 in a boiling water bath for 5 min; 0.1 M sodium acetate; 0.025 M phosphate buffer (pH-7); water; absolute methanol and absolute ether. The samples were then dried overnight in air. Fourier transform infrared spectroscopy (FTIR) analysis was carried out using a Nicolet 6700 FTIR spectrometer coupled with a continuum IR microscope. Operation conditions used a KBr beam splitter and an MCT-A detector (7800–350 cm−1 ). Spectra were obtained in the mid-infrared (4000–400 cm−1 ) region. 32 scans were accumulated for each spectrum, with a spectral resolution of 4 cm−1 .

### Identification of Enzymes Involved in Sporopollenin Biosynthesis

The sporopollenin-related protein sequences of Arabidopsis were downloaded from the Arabidopsis information resource1 (TAIR). The protein sequences of C. protothecoides were then used as a database while the Arabidopsis proteins were used as query sequences for local BLASTP searching to identify homologous protein sequences in C. protothecoides (Wei et al., 2012). We also used CD-Search, a database2 for the functional annotation of proteins (Marchler-Bauer et al., 2005), to identify conserved domains among the protein sequences of the two organisms.

The whole genome sequence has been deposited at DDBJ/EMBL/GenBank under the accession identifier APJO00000000. The sequences for sporopollenin synthesisrelated genes in C. protothecoides have been deposited to the GenBank nucleotide database under the following accession identifiers: KF517419 (Cpr000314.3), KF517420 (Cpr001396.1), KF517421 (Cpr000450.1), KF517422 (Cpr002802.1), KF517423 (Cpr000351.1), KF517424 (Cpr004505.1), KF517425 (Cpr0011 79.1), KF517426 (Cpr002918.1), KF517427 (Cpr001636.6), KF517428 (Cpr004904.1), KF517429 (Cpr001668.1), KF517430 (Cpr004207.2), and KF517431 (Cpr002170.1).

### Real-Time Quantitative PCR

Freshly harvested C. protothecoides cells were quickly frozen and grounded until the liquid nitrogen evaporated. Total RNA was then isolated using Trizol Reagent (Invitrogen, Cat. No. 15596- 026) and genomic DNA was removed by treatment with RNasefree DNaseI (Takara, Cat. No. 2270A). First-strand cDNA was prepared from 4 µg of total RNA using Reverse Transcriptase XL (Takara, Cat. No. 2621) according to the product protocol. PCR amplification was performed using primers designed with Primer Premier 5.0. All primers are listed in Table S2. Actin was used as the internal control. Fluorescence-based real-time PCR reactions were performed in an optical 96-well plate with a Roche LightCycler R 480 II Detection System using SYBR R Green (Invitrogen, Cat. No. 4385618). The 2−11Ct method was used to calculate the changes in relative gene expression measured from the real time quantitative PCR analyses.

### Single Cell Analysis of Asexual Reproduction in *C. Protothecoides* Using a Microfluidic Device

We used a microfluidic system capable of retaining algae cells in microfluidic chambers; this system has been used successfully in yeast for related studies (Tian et al., 2013). This system enables the observation of the asexual reproduction process in single cells throughout their lifespan. The microfluidic device was kindly designed and provided by Chunxiong Luo of Peking University. Images were taken with a Nikon TE2000 time-lapse microscope; bright field images were taken once every 20 min.

### Measurement of Oil Content

Lipid composition and accumulation were analyzed via GC-MS using a slightly modified version of our previously-published protocols (Lu et al., 2010). Wet algal cells treated with or without 2-aminoethanol were suspended in a mixture of 0.5 ml methanol acidified with 3% sulfuric acid and 0.5 ml chloroform containing 2.5 g L−1 capric acid (used as an internal standard to correct transesterification and injection volume errors). The mixture was extracted for different durations of time and then centrifuged at 6000 rpm for 5 min. The supernatant with 3 ml mixture (1.5 ml methanol acidified with 3% sulfuric acid and 1.5 ml chloroform containing 2.5 g L−1 capric acid) was then heated in a sealed tube at 70◦C for 2 h. After cooling, 1 ml of distilled water was added and the sample was vortexed for 20 s. After separation, 1 µl of the lower phase was injected into a gas chromatograph (HP 689-, USA) using a 30 m, 0.32 mm diameter column. Nitrogen was used as the carrier gas at a flow rate of 1 ml min−1 . Measurements started at 80◦C for 1.5 min. The temperature was increased to 140◦C at a rate of 30◦C min−1 . The next step of the method increased the temperature at a rate of 20◦C min−1 to 300◦C and held this temperature for 1 min before the analysis was terminated. The Retention times for the analytes were as follows: 4.4 min for capric acid methyl ester, 7.4 min for hexadecanoic acid, and 8.5 min for octadecanoic acid methyl ester.

### RESULTS

### The Cell Wall of *C. Protothecoides* Includes One Layer Resistant to Cellulose Degrading Enzymes

To study the composition of cell walls, we first stained algal cells with a commonly used cell wall dye, Calcofluor-white, which binds to cellulose and chitin (Hughes and McCully, 2009).

1http://www.arabidopsis.org/

2 http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=cdd

As shown in **Figure 1A**, whole cells stained with Calcofluorwhite showed an intense, light blue fluorescence when irradiated with UV, suggesting that the cell walls of C. protothecoides contain cellulose or chitin, similar to other algae (Popper and Fry, 2003). The enzymatic digestion of algal cell walls has been used for a long time in algal research for the preparation of protoplasts (Aach et al., 1978). Lu et al. described a method for generating protoplasts from C. protothecoides using an a enzyme combination of 2% cellulase and 1% snailase (Lu et al., 2012). We therefore used the same combination of enzymes to treat the Chlorella cells in this study. However, this treatment failed to remove the cell wall (**Figure 1B**). Protoplasts of Chlorella saccharophila, Chlorella ellipsoidea (Braun and Aach, 1975; Yamada and Sakaguchi, 1982), and Chlorella vulgaris (Yang et al., 2015) were obtained using an enzyme mix containing cellulase, hemicellulose, and pectinase. We acquired a number

FIGURE 1 | Enzymatic digestion and Calcofluor-white staining of the cell wall of *C. protothecoides.* The blue fluorescence indicates the presence of a cell wall that contains cellulose or chitin. The red fluorescence is chloroplast autofluorescence. (A) Intact cells; (B) Intact cells after enzymatic treatment; (C) Disrupted cells; (D) Disrupted cells after enzyme treatment. Arrows show cell wall fragments after enzyme treatment.

of commercially available cell wall degrading enzymes, including cellulase, cellulysin, snailase, hemicellulase, pectinase, pectolyase, lysozyme, and zymolase, and applied these, both individually and in various combinations. None of these enzymes could generate protoplasts from C. protothecoides (Table S1).

One of the possibilities for the inability to get protoplasts is that these enzymes are somehow prevented from accessing the cellulose/chitin layer. To test this hypothesis, we partially disrupted the cells in mild to moderately harsh conditions and then stain the sample cells. Pre-disruption of cells with a bead beater in mild conditions retained a round globular structure and showed light blue fluorescence when stained with Calcofluor-white, indicating the presence of a cellulose/chitin layer in the cell wall around these cells (**Figure 1C**). Interestingly, following treatment with an enzyme combination of 2% cellulase and 1% snailase, most cells failed exhibit fluorescence upon Calcofluor-white staining, suggesting the loss of cellulose/chitin layer around these cells (**Figure 1D**). These results indicate that cellulose/chitin is one of the major components of the C. protothecoides cell wall and is protected from enzymatic degradation by a hitherto unknown mechanism.

### Identification of Sporopollenin in the Cell Wall of *C. Protothecoides*

In a quest to understand what prevents the cell wall being subjected to enzymatic degradation, we examined the ultrastructure of the C. protothecoides cell wall using TEM. Isolated cell walls were subjected to TEM analysis, at different magnitudes, both for cells cultured heterotrophically (**Figure 2**) and for cell cultured autotrophically (Figure S1). As shown in **Figures 2A–F**, the cell wall of heterotrophic C. protothecoides consists of two morphologically distinct components that are referred to henceforth as the "inner" and "outer" parts of the wall. The inner part is similar to the cell wall of many plants and presumably consists of polysaccharides and proteins. The inner wall of heterotrophic algae (**Figures 2C–F**) is about 80–100 nm in thickness; wider than the inner part of the walls of autotrophic algae cells, which is about 50–60 nm (Figures S1C–F). A mixture of 2% cellulase and 1% snailase was about to completely digest the inner wall of both heterotrophic (**Figures 2G–I**) and autotrophic algae (Figures S1G–I). This observation suggested that the inner wall is probably composed of cellulose and protein, similar to the cell walls of other plants or algae (Lora et al., 2014). It is known that the polysaccharide components in the cell wall can support the structure and keep cells in a relatively constant circular state, so it was not surprising that the loss of polysaccharides following enzymatic treatment resulted in curved cell wall fragments (**Figure 2G**, Figure S1G).

The outer layer of both autotrophic and heterotrophic algae displayed a trilaminar composition and was about 10 nm in thickness (**Figure 2C**). It was sandwich-like in structure and included an electron-lucent center layer and two electrondense zones on either side of this layer. Interestingly, the outer layer was resistant to enzymatic digestion, indicating that it was probably not made of cellulose or other polysaccharides. Previous studies have suggested that the outer layer of pollen from many plants is resistant to enzymatic digestion and is composed of sporopollenin, a material that can be dissolved in oxidizing solutions, fused potassium hydroxide, or 2-aminoethanol (Southworth, 1974; Domínguez et al., 1999). Therefore, we treated the isolated cell walls with 2 aminoethanol and found that the outer layer became smeared although the inner layer did not undergo significant changes (**Figures 3A–C**). This experiment that there is an outer layer of the C. protothecoides cell wall that is possibly composed of sporopollenin.

To evaluate whether or not the newly-identified layer is indeed made of sporopollenin, two additional assays were performed. First, the algal cells were subjected to acetolysis, a process to which sporopollenin is known to be resistant (Heslop-Harrison, 1969). As a control, yeast cells were treated under the same conditions. As shown in **Figure 3D**, the yeast cells were completely dissolved following acetolysis, resulting in a transparent solution. With the C. protothecoides samples, however, a large amount of insoluble material remained after treatment. This result is similar findings of Good and Chapman, who identified sporopollenin-containing plants (Good and Chapman, 1978). We next analyzed the composition of the outer layer directly using infrared spectroscopy. We observed two large peaks around the 3000 cm−1 wavelength, and noted a slight variation from 1400 to 1500 cm−1 (**Figure 3E**, Figure S2). This spectral data matches the previously-reported spectra of pollen as well as the outer layer of the cell walls of other sporopollenincontaining algae (Watson et al., 2007). Taken together, these results strongly support the assertion that sporopollenin is present in the outer layer of the cell walls of C. protothecoides.

### Identification of Genes Likely Involved in Sporopollenin Biosynthesis in the Genome of *C. Protothecoides*

Genomic analysis of sporopollenin formation in higher plants has indicated that sporopollenin shares a common synthesis pathway, and a schematic model of sporopollenin formation has been proposed (Ma, 2005; Blackmore et al., 2007). We recently published a draft genome of C. protothecoides (Gao et al., 2014). This genome enabled us to identify genes in this alga that are homologous to enzymes known or thought to function in key roles in the sporopollenin synthesis pathway. Using Arabidopsis homologs as reference, we found that almost all of the genes known to be associated with sporopollenin formation could be identified in the C. Protothecoides genome (**Table 1**); further analysis resulted in the identification of conserved domains among these enzymes. The key genes for sporopollenin formation are visually summarized as three steps in C. protothecoides, as shown in **Figure 4A**. First, the formation of a callose wall that separates individual daughter cells; this is likely catalyzed by Cpr000314.3 and Cpr001396.1, which are the homologs, respectively, of CaLS5 and KNS2 in Arabidopsis. Pollen wall development is subsequently initiated through primexine formation around distinct daughter cells. Previous studies in plants have shown that that primexine functions as a sporopollenin receptor; four enzymes, DEX1, MS1,

MCW, mother cell wall; Triangles show the inner layer of the cell wall; Arrows show the outer layer of the cell wall.

NEF1, and RPG1 are required for sporopollenin deposition and polymerization (Gabarayeva et al., 2009). Similarly, Cpr000450.1, Cpr002802.1, Cpr000351.1, and Cpr004505.1, which correspond, respectively, to the genes that encode these enzymes, were identified in the genome of C. protothecoides. The third step is the synthesis, secretion, and translocation of sporopollenin precursors, a process that is dependent on the synthesis of free fatty acids and their derivatives (Meuter-Gerhards et al., 1999, **Figure 4A**). Multiple enzymes are known to function in these metabolic pathways, and most of these were found in the C. protothecoides genome (**Table 1**). Genetic studies in Arabidopsis and other organisms have shown that mutations in the aforementioned genes, including, for example CaLS5 and KNS2, lead to defects in primexine formation (Ariizumi et al., 2004; Dobritsa et al., 2010; Li et al., 2010). The identification of similar genes in the genome of C. protothecoides establishes a putative molecular basis for the existence of sporopollenin in this alga.

heterotrophic algae, and autotrophic algae. Samples before Acetolysis treatment were on the top and after treatment were at the bottom. (E) FTIR of the cell wall of *C. protothecoides*. SC, Saccharomyces cerevisiae; HCP, Heterotrophic *C. protothecoides*; ACP, Autotrophic *C. protothecoides*. Triangles show the inner layer of the cell wall.

To further verify the existence of above identified genes related to sporopollenin biosynthesis, primers were designed according to the sequences of the conserved domains found in C. protothecoides genome; PCR products of the expected sizes were amplified from C. protothecoides genomic DNA (Figure S3). Additionally, real time PCR was used to compare the expression of sporopollenin-synthesis genes in autotrophic and heterotrophic cells. As shown in **Figure 4B**, transcripts of all of these genes could be detected under both conditions. Of note, as compared to the cells growing in the autotrophic condition, the heterotrophic cells tended to have higher expression of these genes. It is possible that cells growing in the heterotrophic condition undergo relatively more rapid replication and that relatively higher abundances of enzymes are therefore required to satisfy growth and division demands.

### The Sporopollenin Layer Exists Throughout the Entire Asexual Life Cycle and Forms an Obstacle for Oil Extraction

The continuous tracking of algal cells in a microfluidic device makes it possible to study the cell division dynamics of single cell with high temporal resolution. We monitored a time course of C. protothecoides cell division and observed that protoplasts divide into two, four, eight, and so on, within the mother cell wall (**Figure 4C**). When the mother cell wall ruptures, the daughter cells emerge from the autosporangium; this is a common mode of propagation for unicellular green algae. We present the entire process of the cell cycle arranged into a movie (Movie S1). Algae growing in both autotrophic and heterotrophic media showed the same division pattern (Figure S4). This propagation mechanism immediately suggests that an intact cell wall will be formed for each daughter cell within a mother cell; this was confirmed by transmission electron microscopy analysis (**Figure 4C**). This also suggests that it is not possible to obtain cells without this sporopollenin layer via targeted collection of mother cells. Therefore, an additional method to remove the sporopollenin layer might be required in order to improve the efficiency for both transformation and oil extraction.

To test this hypothesis, C. protothecoides cells were pre-treated with 2-aminoethanol, which was able to dissolve sporopollenin in the aforementioned experiments in our study and in previouslyreported experiments (Southworth, 1974; Domínguez et al., 1999, **Figure 3A**). As expected, we found that the oil yield

*CWS, Callase wall Synthesis pathway; PS, Primexine Synthesis; SST, Sporopollenin synthesis secretion and translocation.*

was significantly improved after 2-aminoethanol treatment as compared the no treatment control cells (**Figure 4D**). As we reported before (Xiao et al., 2015), oil extraction from C. protothecoides is one of the most energy-consuming steps in large scale biofuel production procedures. Our results suggest that a potential solution to help reduce the cost of this step would be the pre-treatment of cells with 2-aminoethanol.

### DISCUSSION

Although, C. protothecoides is regarding a good candidate species for biofuel production (Heredia-Arroyo et al., 2010), its cells are notorious their hard-to-disrupt cell wall. This results in an increased cost for oil extraction from these algal cells. According to previous oil production studies, a pre-treatment process is always required to enhance the efficiency of lipid extraction (Halim et al., 2012). Currently, however, no mature pretreatment methods can be directly applied to C. protothecoides to extract oil at an industrial scale. Actually, when typical methods are used, the recovery rate of lipids from C. protothecoides is not as high as that of other organisms (Shen et al., 2009). It has been a long-standing mystery as to why this particular alga is so resistant to mechanical stress. Several attempts to genetically transform C. protothecoides have been unsuccessful. Common methods such as the use of polyethylene glycerol following spheroplast generation, which are used to transform protoplasts of other yeast and fungal cells (Kindle, 1990), do not work for C. protothecoides. And direct enzymatic digestion using cellulase, snailase or other cell wall-degrading enzymes cannot remove the C. protothecoides cell wall (**Figures 1B**, **2B,D**). The identification of sporopollenin as a major component of the cell wall provides a reasonable explanation to these questions.

Knowing the composition of cell wall should greatly advance the use of C. protothecoides as model system for biofuel production. For example, additional pre-treatment steps can be adopted to remove sporopollenin prior to oil extraction, likely leading to increased oil yields. In addition, the identification of the sporopollenin layer promises to facilitate the development of new methods for the genetic transformation of C. protothecoides. A previous study suggested that microprojectile bombardment could be used to deliver DNA into tobacco pollen to study the expression of pollen-expressed genes (Twell et al., 1989). Given the superficially similar cell wall structure between C. protothecoides cells and pollen, it is conceivable that a similar method might also work to transform this alga. Furthermore, screens can be carried out to find mutants in which the synthesis of sporopollenin is blocked. Alternatively, some specific chemicals may be used to inhibit sporopollenin synthesis. These sporopollenin deficient strains may become susceptible and easy to accept foreign DNA.

Recently, an elegant study that systematically treated various Chlorella strains with a large number of commercially available

enzymes found that Chlorella is most sensitive to chitinase and lysozymes (Gerken et al., 2013). We performed a similar series of enzymatic treatments with C. protothecoides. Unfortunately, none of the enzymes we tested were capable of removing the cell wall of C. protothecoides. This frustrating result actually lead us to carefully examine its cell wall with electron microscopy, leading to our discovery of the sporopollenin layer (**Figure 2C**).

Sporopollenin is considered to be one of the toughest materials in nature and is known to be resistant to many kinds of chemical and biological attacks. It has been found in Characium terrestre, Enallax coelastroides, and Scotiella chlorelloidea as a high efficient barrier against physical or chemical damage and has been extremely important in the evolution of terrestrial plants (Xiong et al., 1997). However, only a few green algae species have been reported to contain sporopollenin in their cell walls (Northcote et al., 1958; Krienitz et al., 1999). The function of sporopollenin in microalgae has been proposed to be associated with resistance to environmental stress such as UV irradiation, desiccation, microbiological attack, and so on (Strohl et al., 1977).

Similarly, the presence of sporopollenin in the cell wall may help C. protothecoides to withstand various environmental stresses.

### AUTHOR CONTRIBUTIONS

QW, JD conceived research; JD, XH designed experiments; XH performed experiments and analyzed the data; all authors wrote, commented on, and approved the contents of the manuscript.

### ACKNOWLEDGMENTS

We are grateful to Dr. Vanita Uppada for critical reading of this manuscript, and grateful to Zhouqing Luo and members

### REFERENCES

of the Wu lab for helpful discussions. We thank the Center of Biomedical Analysis at Tsinghua University for help with the transmission electron microscopy service. This work was supported by the National Natural Science Foundation of China (NSFC) 41030210, the Ministry of Science and Technology (MOST) projects 2011BAD14B05 and 2011CB808804 to QW, and by the Tsinghua University Initiative Scientific Research Program<PHONE_NUMBER>6 to JD.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2016.01047

further evidence for the participation of self-assembly processes. Rev. Palaeobot. Palynol. 156, 211–232. doi: 10.1016/j.revpalbo.2008.12.001

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 He, Dai and Wu. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Photoconversion and Fluorescence Properties of a Red/Green-Type Cyanobacteriochrome AM1\_C0023g2 That Binds Not Only Phycocyanobilin But Also Biliverdin

Keiji Fushimi1 , Takahiro Nakajima2 , Yuki Aono2 , Tatsuro Yamamoto1 , Ni-Ni-Win2 , Masahiko Ikeuchi2,3, Moritoshi Sato2 and Rei Narikawa1 \*

1 Department of Biological Science, Faculty of Science, Shizuoka University, Shizuoka, Japan, 2 Graduate School of Arts and Sciences, University of Tokyo, Tokyo, Japan, 3 Core Research for Evolutional Science and Technology, Japan Science and Technology Agency, Saitama, Japan

#### Edited by:

Takashi Osanai, Meiji University, Japan

#### Reviewed by:

Youn-Il Park, Chungnam National University, South Korea Andreas Möglich, Universität Bayreuth, Germany Scheer Hugo, Universität München, Germany

> \*Correspondence: Rei Narikawa<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology

> Received: 28 January 2016 Accepted: 11 April 2016 Published: 26 April 2016

#### Citation:

Fushimi K, Nakajima T, Aono Y, Yamamoto T, Ni-Ni-Win, Ikeuchi M, Sato M and Narikawa R (2016) Photoconversion and Fluorescence Properties of a Red/Green-Type Cyanobacteriochrome AM1\_C0023g2 That Binds Not Only Phycocyanobilin But Also Biliverdin. Front. Microbiol. 7:588. doi: 10.3389/fmicb.2016.00588 Cyanobacteriochromes (CBCRs) are distantly related to the red/far-red responsive phytochromes. Red/green-type CBCRs are widely distributed among various cyanobacteria. The red/green-type CBCRs covalently bind phycocyanobilin (PCB) and show red/green reversible photoconversion. Recent studies revealed that some red/green-type CBCRs from chlorophyll d-bearing cyanobacterium Acaryochloris marina covalently bind not only PCB but also biliverdin (BV). The BV-binding CBCRs show far-red/orange reversible photoconversion. Here, we identified another CBCR (AM1\_C0023g2) from A. marina that also covalently binds not only PCB but also BV with high binding efficiencies, although BV chromophore is unstable in the presence of urea. Replacement of Ser334 with Gly resulted in significant improvement in the yield of the BV-binding holoprotein, thereby ensuring that the mutant protein is a fine platform for future development of optogenetic switches. We also succeeded in detecting nearinfrared fluorescence from mammalian cells harboring PCB-binding AM1\_C0023g2 whose fluorescence quantum yield is 3.0%. Here the PCB-binding holoprotein is shown as a platform for future development of fluorescent probes.

Keywords: optogenetics, linear tetrapyrrole, GFP, near-infrared fluorescence, live cell imaging

## INTRODUCTION

Phytochromes and cyanobacteriochromes (CBCRs) are photoreceptors that form a large superfamily with a linear tetrapyrrole-binding GAF (cGMP-phosphodiesterase/adenylate cyclase/FhlA) domain (Ikeuchi and Ishizuka, 2008; Anders and Essen, 2015). Some CBCRs are known to be involved in regulation of light acclimation processes such as phototaxis (Yoshihara et al., 2000; Narikawa et al., 2011; Song et al., 2011; Savakis et al., 2012; Campbell et al., 2015), chromatic acclimation (Kehoe and Grossman, 1996; Hirose et al., 2010) and light-dependent cell aggregation (Enomoto et al., 2014, 2015). Only GAF domain of CBCRs is enough for chromophore ligation and photoconversion, although additional PAS (Per/Arnt/Sim) and PHY

(phytochrome-specific) domains in phytochromes are necessary. CBCRs have been roughly categorized into two types according to the chromophore they contain in the thermostable state: Phycoviolobilin (PVB) and PCB. In both cases, PCB is initially incorporated into the GAF domain and Cys residue within the GAF domain covalently ligates to C31 of the chromophore.  These CBCRs commonly show light-induced Z/E isomerization of a double bond between rings C and D, followed by various structural changes such as reversible protochromic cycle and reversible Cys-adduct formation (Rockwell et al., 2008; Burgie et al., 2013; Hirose et al., 2013; Narikawa et al., 2013, 2014).

 The red/greentype CBCRs show reversible photoconversion between a redabsorbing form (Pr) with 15Z-PCB and a green-absorbing form (Pg) with 15E-PCB. Structure of Pr form provides direct insights into chromophore–protein interaction (Narikawa et al., 2013). Detailed spectroscopic analyses based on this structure have revealed their photoconversion mechanism in which excited state destabilization and ring D distortion are suggested to occur to form blue-shifted Pg form upon red-light irradiation (Rockwell et al., 2014; Song et al., 2015a).

Recently, it has been revealed that red/green-type CBCRs (AM1\_1557g2 and AM1\_1870g3) derived from the chlorophyll d-bearing cyanobacterium Acaryochloris marina covalently bind not only PCB but also biliverdin (BV; Narikawa et al., 2015a,b). BV-binding ones show reversible photoconversion between far red-absorbing (Pfr) form and orange-absorbing (Po) form, whereas PCB-binding ones show normal red/green reversible photoconversion. Site-directed mutagenesis suggests that a Cys residue within the GAF domain covalently ligates not only to PCB but also to BV. BV is present in most organisms including mammals and far-red light can penetrate into deep tissues, with a potential as optogenetic and bioimaging tools (Ziegler and Möglich, 2015).

Here, we report another GAF domain (second GAF domain of AM1\_C0023 called AM1\_C0023g2) from A. marina that covalently binds not only PCB but also BV. Replacement of Ser334 with Gly resulted in significant improvement in yield of the BV-binding holoprotein. Further, we detected near-infrared fluorescence from mammalian cells harboring AM1\_C0023g2-PCB whose fluorescence quantum yield is 3.0%.

### MATERIALS AND METHODS

### Plasmid Construction

The nucleotide sequence of AM1\_C0023g2 was cloned into pET28a (Novagen) using In-fusion HD Cloning kit (TaKaRa) as described previously (Narikawa et al., 2015b). AM1\_C0023g2 sequence was amplified by polymerase chain reaction (PCR) with a specific primer set (50 - CGCGGCAGCCATATGAATATTTCCGAGATTATT-30 , 50 -CT CGAATTCGGATCCTCAAGCTTCTGCTTTGTTTTT-30 ) and PrimeSTAR Max DNA polymerase (TaKaRa). The inserted sequence was confirmed by sequencing with an ABI310 genetic analyzer. Replacement of AM1\_C0023g2 Ser334 with Gly (denoted S334G) was performed using a specific primer set (50 -CAACAAGGATATACAGATTGTCATCTA-30 , TGTATATCCTTGTTGATAAATGTCAGC) and PrimeSTAR max DNA polymerase as described previously.

To construct GFP-fused AM1\_C0023g2 and AM1\_1557g2, the nucleotide sequences of GFP, AM1\_C0023g2 and AM1\_1557g2 were amplified by PCR with specific primer sets and Pyrobest DNA polymerase (TaKaRa). The following primer sets were used to introduce restriction enzyme sites, a flexible

FIGURE 2 | Photoconversion of AM1\_C0023g2-PCB and AM1\_C0023g2-BV. (A) Absorption spectra of Pr (magenta) and Pg (blue) forms of AM1\_C0023g2-PCB. (B) Absorption spectra of Pfr (magenta) and Po (blue) forms of AM1\_C0023g2-BV. (C) Difference spectra of PCB- (blue) and BV-binding (magenta) AM1\_C0023g2 before and after photoconversion.

peptide linker sequence, and Kozak sequence: for GFP, 50 - ATGCAAGCTTGCCACCATGGTGAGCAAGGGCGAG-30 and 5 0 -GCATCTCGAGACCTCCGCTACCGCCCTTGTACAGCTCG TC-30 ; for AM1\_C0023g2 and AM1\_1557g2, 50 -ATGC CTCGAGAGCGGCCTGGTGCCGCGC-30 and 50 -GAGCTCG AATTCGGATCC-30 . All sequences were confirmed by sequencing with an ABI 310 genetic analyzer. These constructs were cloned into a mammalian expression vector pcDNA 3.1 (+) (Invitrogen) using the restriction enzyme sites. All the mammalian expression plasmids were purified using QIAGEN plasmid kit (Qiagen).

### Expression, Purification, and SDS-PAGE

His-tagged AM1\_C0023g2 was expressed in both BV- and PCBproducing Escherichia coli (C41 harboring pKT270 and pKT271, respectively). The His-tagged proteins were isolated by Niaffinity chromatography as described previously (Narikawa et al., 2015b). The purified proteins were subjected to SDS-PAGE, followed by in-gel fluorescent assay and Coomassie Brilliant Blue staining as described previously (Narikawa et al., 2015b). Fluorescence was visualized through a 600 nm long path filter upon excitation with wavelength of blue (λmax = 470 nm) and green light (λmax = 530 nm) through a 562 nm short path filter (WSE-6100 LuminoGraph, WSE-5500 VariRays; ATTO).

### Estimation of Binding Efficiencies

Protein concentration was determined by standard Bradford method (Bio-Rad). We determined extinction coefficients of free PCB (Santa Cruz Biotechnology Inc.) and free BV (Frontier Scientific) under both acidic urea and 1% SDS conditions. Extinction coefficients of the free PCB and BV under the acidic urea condition were calculated to be a little less than 30000, that is comparable to that reported previously (Glazer and Fang, 1973). On the other hand, extinction coefficients of the free PCB and BV under the 1% SDS condition were calculated to be around 17000. Although extinction coefficients of chromophores under the 1% SDS condition tend to be lower than those under the acid urea condition, PCB and BV behave similarly under both conditions. Thus, concentration of PCB bound to proteins was determined by absorbance at 666 nm when denatured with 8 M urea (pH2.0). Similarly that of BV bound to proteins was determined by absorbance at 646 nm when denatured with 1% SDS.

### In Vitro Reconstitution

S334G protein that is a mutant of AM1\_C0023g2 was used for in vitro reconstitution analysis because of its high yield of in vivo reconstitution system (see **Figure 4**). Free BV and the apo-S334G protein were mixed in roughly equimolar amounts and incubated for 30 min at 37◦C. After removal of unbound BV by spin column, the sample was subjected to spectral analyses.

## Spectroscopy

Ultraviolet and visible absorption spectra were recorded with a Shimadzu UV-2600 spectrophotometer. Monochromic light of various wavelengths for photoconversion was generated using a variable wavelength light source (Opto-Spectrum Generator, Hamamatsu Photonics, Inc.). Acid denaturation of the proteins was performed with 8 M urea, pH 2.0 under the dark condition. Fluorescence spectra were recorded with a StellarNet SILVER-Nova spectrometer (StellerNet, Inc.). Fluorescence quantum yields were measured with Quantaurus-QY (Hamamatsu Photonics, Inc.).

### Imaging of GFP-fused AM1\_C0023g2 and AM1\_1557g2 in Mammalian Cells

Human cervical carcinoma (HeLa) cells (ATCC, CCL-2) were cultured in Eagle's minimum essential medium (Sigma) supplemented with 2 mM L-glutamine (Gibco), 10% fetal bovine serum (Gibco), 100 U/mL penicillin and 100 µg/mL streptomycin (Gibco) at 37◦C in 5% (v/v) CO2. The day before transfection, HeLa cells were plated onto 35-mm glass-based dishes (Iwaki). Transfection was performed using lipofectamine 3000 (Life Technologies) according to the manufacturer's instructions. Forty-eight hours after transfection, the cells were washed with Hank's balanced salt solution (HBSS; Gibco) and replaced with HBSS containing either 20 µM PCB (Frontier Scientific) diluted from a stock solution of 20 mM in dimethyl sulfoxide (DMSO; Wako) or 0.1% (v/v) DMSO as the vehicle control. After incubation for 4 h, the cells were washed and replaced with HBSS, and imaged using a Zeiss LSM 710 confocal microscope equipped with a Plan-Apochromat 63×/1.4 oil immersion objective (Carl Zeiss), a 488 nm Ar laser, and a 633 nm He/Ne laser. Green fluorescence of GFP was detected at 493–538 nm upon excitation with the 488 nm laser (6.5 µW measured at the back aperture of the objective lens). Nearinfrared fluorescence of CBCR was detected at 638–740 nm upon excitation with the 633 nm laser (67 µW measured at the back aperture of the objective lens). Data analysis was performed using ZEN 2009 (Carl Zeiss).

## RESULTS AND DISCUSSION

### AM1\_C0023g2 Isolated from PCB-Producing and BV-Producing E. coli

We found that AM1\_C0023 is closely related in-paralog of AM1\_1557. AM1\_C0023 is 71% identical to and has a domain architecture same as AM1\_1557 (**Figure 1A**). Second GAF domain of AM1\_C0023 (AM1\_C0023g2) is 84% identical to that of AM1\_1557 (AM1\_1557g2). Residues highly conserved among red/green-type CBCRs and important for chromophore ligation are also conserved in AM1\_C0023g2.

AM1\_C0023g2 was expressed in both PCB- and BVproducing E. coli and purified by using nickel-affinity column chromatography. In-gel fluorescence analysis revealed that AM1\_C0023g2 covalently binds not only PCB but also BV like AM1\_1557g2 (**Figure 1B**). PCB-binding AM1\_C0023g2 (AM1\_C0023g2-PCB) showed reversible photoconversion between red-absorbing Pr form at 650 nm and green-absorbing

Pg form at 539 nm (**Figure 2A**), whereas BV-binding AM1\_C0023g2 (AM1\_C0023g2-BV) showed reversible photoconversion between far red-absorbing Pfr form at 699 nm and orange-absorbing Po form at 618 nm (**Figure 2B**). These spectral features are almost same as those of AM1\_1557g2 (Narikawa et al., 2015b). Pfr-minus-Po difference spectrum of AM1\_C0023g2-BV possesses positive peaks at 700 and 377 nm and negative peaks at 601 and 284 nm, whereas Pr-minus-Pg difference spectrum of AM1\_C0023g2-PCB possesses positive peaks at 650, 352, and 301 nm and negative peaks at 536 and 270 nm (**Figure 2C**).

AM1\_C0023-BV Pfr showed dark reversion to Po (**Figure 3A**), whereas dark reversion of AM1\_C0023-PCB Pr was hardly detected (**Figure 3B**). Half-lives for the AM1\_C0023-BV during

dark reversion measured at 15, 20, and 25◦C were 1356, 478, and 180 min, respectively (**Figure 3B**). Slower dark reversion was observed under lower temperature. Hardly detectable and relatively fast dark reversions of PCB- and BV-binding ones, respectively, are also consistent with those found for AM1\_1557g2 and AM1\_1870g3 (Narikawa et al., 2015a,b).

### Site-Directed Mutagenesis of AM1\_C0023g2

In the process of site-directed mutagenesis analysis to obtain proteins with higher fluorescence quantum yields, we incidentally obtained a unique mutant protein, S334G in which Ser334 was replaced with Gly. This replacement resulted in significant improvement in expression yield of BV-binding holoproteins in E. coli (**Figure 4A**). Colonies were picked up and grown in small-scale culture. Cell pellet of E. coli expressing only the wild type protein showed brown color typical of normal E. coli. Cell pellet of E. coli expressing both the wild type protein and the heme oxygenase showed pale green color. On the other hand, cell pellet of E. coli expressing both the S334G protein and the heme oxygenase showed deep green color. The BVbinding S334G (S334G-BV) showed reversible photoconversion between Pfr at 699 nm and Po at 611 nm (**Figure 4B**). Its Po form is about 7 nm blue-shifted than that of the wild type. Binding efficiency to BV was calculated to be 1.25-fold higher than that of the wild type protein (see Evaluation of Chromophore Binding Efficiencies). In total, judging from the Pfr absorption peak, recovery of the S334G-BV was about fivefold higher than that of the BV-binding wild type from the same culture volume. Although Gly317 of AnPixJg2 corresponding to Ser334 of AM1\_C0023g2 does not directly interact with PCB, Gly317 is placed 4 residues upstream of Cys321 that covalently ligates to PCB (Narikawa et al., 2013). This replacement may indirectly affect chromophore-binding pocket to facilitate protein expression and chromophore incorporation. Apart from the underlying mechanism, we consider that S334G-BV is a good platform for further development of optogenetic switches. Slow dark recovery kinetics and almost full photoconversion (90% estimated from the absorption spectra) are favorable for longterm and acute regulation.

### Chromophore Species and Configuration

Acid-denatured spectra proved that AM1\_C0023g2 isolated from PCB-producing E. coli undoubtedly bound PCB (**Figure 5**). Absorption maximum of denatured Pr and Pg were observed at 665 and 603 nm, respectively (**Figures 5A,B**). Irradiation of denatured Pg with white light resulted in red shift of the absorbance that is almost identical to that of denatured Pr, whereas irradiation of denatured Pr with white light resulted in no significant change (**Figures 5A,B**). Pr-minus-Pg difference spectrum fits to that during photoconversion of denatured Pg (**Figure 5C**). These results clearly show that Pr and Pg bound 15Z- and 15E-PCB, respectively.

Acid-denatured spectra of AM1\_C0023g2 isolated from BVproducing E. coli showed completely different spectra (data not shown) compared with previously reported data (Narikawa

FIGURE 6 | Acid denaturation of AM1\_C0023g2-BV. (A) Absorption spectra of acid-denatured AM1\_C0023g2-BV Pfr during dark and light processes. (B) Absorption spectra of acid-denatured AM1\_C0023g2-BV Po during dark and light processes. (C) Difference spectrum during dark and light processes of denatured Pfr (magenta), during dark and light processes of denatured Po, and of Pfr-minus-Po just after denaturation (green). (D) Normalized absorption spectra and difference spectrum of acid-denatured AM1\_C0023g2-BV Pfr and Po after dark and light processes.

et al., 2015b). In order to monitor initial spectral change after denaturation, we denatured samples under 4◦C and their absorption spectra were measured at 15◦C at 0, 1, 3, 5, and 10 min. Subsequently absorption spectra during white light illumination (1, 3, 5, and 10 min) were recorded. Absorption maximum just after denaturation of Pfr form was observed at ∼700 nm (**Figure 6A**), which corresponded to 15Z-BV previously reported (Narikawa et al., 2015b). Absorption maximum after denaturation of Po form was observed at ∼660 nm (**Figure 6B**) with a red-shift compared with 15E-BV as reported earlier. Because photoconversion from Pfr to Po is incomplete (**Figure 2B**), the residual Pfr may affect the absorption peak of denatured Po resulting in red shift to ∼660 nm.

The ∼700 nm peak observed for Pfr (denatured) decreased during dark and light processes and final absorption spectrum was at ∼665 nm (**Figure 6A**). Difference spectrum during these processes showed a negative peak at ∼714 nm (**Figure 6C**, Pfr W10 – Pfr D0). From these results, we hypothesized that 15Z-BV just after denaturation rapidly converts to an unknown state (X) at 665 nm. On the other hand, absorption after Po denaturation slightly decreased under dark process and largely decreased under light process (**Figure 6B**). The decrease under dark process may correspond to conversion from the residual Pfr to the unknown state "X". The decrease under light process may correspond to conversion from Po to "X" via Pfr. Based on the above-mentioned hypothesis, in the case of both Pfr denaturation and Po denaturation, their final products after dark and light processes should be composed of only "X". Indeed, absorption spectra of both denatured Pfr and Po after 10 min white light illumination are at 665 nm and almost same (**Figure 6D**), which strongly supports this hypothesis.

Pfr-minus-Po difference spectrum just after denaturation showed a positive peak around ∼714 nm and a negative peak around ∼614 nm (**Figure 6C**, Pfr D0 – Po D0). These wavelengths well correspond to those obtained from the difference spectrum of denatured AM1\_1557g2 (Narikawa et al., 2015b). Thus, the positive peak around 714 nm corresponds to 15Z-BV, whereas the negative peak around 614 nm corresponds to 15E-BV. However, a negative component around 745 nm was detected. Based on the above-mentioned hypothesis in which the denatured 15Z-BV rapidly converts into "X", 15E-BV from denatured Po should be more abundant than 15Z-BV from denatured Pfr. In such situation, absorption tail of 15E-BV around 745 nm should be larger than that of 15Z-BV, and the difference spectrum should show the minus component around 745 nm.

Difference spectrum during light and dark processes of denatured Po showed negative peaks at ∼714 and ∼614 nm (**Figure 6C**, Po W10 – Po D0). These wavelengths also correspond to 15Z-BV and 15E-BV, respectively. Major negative peak at ∼614 nm is likely to correspond to conversion from 15E-BV to "X" via 15Z-BV, whereas minor negative peak at ∼714 nm is likely to correspond to conversion from 15Z-BV originated in the residual Pfr to "X".

FIGURE 7 | Denaturation with 1% SDS and in vitro reconstitution. (A) Absorption spectrum of AM1\_C0023g2-BV Pfr denatured with 1% SDS. (B) Absorption spectrum of AM1\_1557g2-BV Pfr denatured with 1% SDS. (C) Absorption spectra of the Pfr and Po forms of S334G reconstituted with BV in vitro. Photoconversion was repeated twice. First-round Po: Solid blue line, first-round Pfr: Solid magenta line, second-round Po: Dotted cyan line, second-round Pfr: Dotted pink line. Inset: SDS-PAGE of in vitro reconstituted S334G. Left, CBB-stained gel. Right, in-gel fluorescence detection. (D) Pfr-minus-Po difference spectra of S334G reconstituted in vitro and in vivo. In vitro reconstitution: blue (first-round) and magenta (second-round) lines, in vivo reconstitution: black line.

Taking the possibility into consideration that the BV chromophore bound to AM1\_C0023g2 is unstable under acidic condition, we denatured the AM1\_C0023g2-BV under neutral urea condition. However, the denatured chromophore was also unstable and its absorption was rapidly bleached (data not shown). The result suggests that urea is crucial for the chromophore instability. Thus, we denatured the AM1\_C0023g2-BV Pfr under 1% SDS as previously described (Ishizuka et al., 2007). As a result, its absorption maxima was observed at ∼646 nm and almost same as that of AM1\_1557g2-BV Pfr (**Figures 7A,B**), indicating that their chromophores are both BV. To further confirm the result, we performed in vitro reconstitution analysis using S334G protein and BV chromophore. Although reconstitution efficiency was low, BV covalently bound to S334G was detected by ingel fluorescence analysis (**Figure 7C**, inset). Far-red/orange reversible photoconversion was observed (**Figure 7C**) and is almost same as that observed for in vivo reconstituted one (**Figure 7D**). As a conclusion, the chromophore of AM1\_C0023g2 expressed in BV-producing E. coli is indeed BV.

FIGURE 8 | Fluorescence spectra of AM1\_C0023g2-BV Pfr (magenta) and AM1\_C0023g2-PCB Pr (blue).

## BV Chromophore Instability in the Presence of Urea

Although AM1\_C0023g2 binds BV as well as the other BVbinding CBCRs and bacteriophytochromes, BV derived from only AM1\_C0023g2 showed unstable nature in the presence of urea. Very recently, Shcherbakova et al. (2015) obtained a fluorescent protein that consists of PAS and GAF domain originally derived from a bacteriophytochrome, RpBphP1 (Shcherbakova et al., 2015). The fluorescent protein, Bph1-FP, covalently binds BV via Cys residue within the GAF domain. The crystal structure revealed that covalent-bonding site of Bph1-FP to BV is heterogeneous, namely C31 or C32 . This means that a Cys residue within the GAF domain has a potential to ligate to both C31 and C32 . In this context, we hypothesized that the Cys residue of AM1\_C0023g2 ligates to C31 and the resultant adduct between the Cys residue and C31 of BV may affect stability of the denatured chromophore in the presence of urea, whereas Cys residues of AM1\_1557g2 and AM1\_1870g3 are likely to ligate to C32 of BV as well as normal bacteriophytochromes and the resultant chromophore may be stable under the same condition. Just after denaturation of Pfr form, a component

with PCB. Left column, GFP-fused AM1\_1557g2 with PCB; right column, GFP-fused AM1\_C0023g2 with PCB; green images, GFP fluorescence; red images, fluorescence of CBCR with PCB. Scale bars, 20 µm. (C) Relative intensity of fluorescence of CBCR with PCB normalized to GFP fluorescence. Error bars represent standard deviations (n = 16, 16, 16, and 17 for AM1\_1557g2 with vehicle, AM1\_1557g2 with PCB, AM1\_C0023g2 with vehicle and AM1\_C0023g2 with PCB, respectively).

corresponding to BV and at 700 nm rapidly converts into a component at 665 nm. This characteristic in absorbing around 665 nm is somehow similar to that of PCB. In this context, BV bound to AM1\_C0023g2 may break down into PCB-like chromophore in the presence of urea. Urea may affect conjugated system of BV bound to AM1\_C0023g2. In order to verify these hypotheses, we are now trying to crystalize both AM1\_1557g2- BV and AM1\_C0023g2-BV.

### Evaluation of Chromophore Binding Efficiencies

We calculated binding efficiencies of AM1\_C0023g2 to PCB and BV at ∼80 and ∼70%, respectively. Based on these values and in-gel fluorescence analysis (**Figure 1B**), we also estimated the binding efficiencies of AM1\_1557g2 to PCB and BV at ∼50 and ∼40%, respectively. The binding efficiencies of AM1\_C0023g2 to both PCB and BV are higher than those of AM1\_1557g2. The binding efficiency of S334G to BV was calculated to be almost 100%, which is better than that of the wild type protein (∼70%).

Binding efficiency of AM1\_C0023g2 to PCB is comparable to or slightly higher than that to BV. In this context, it is intriguing that no BV-binding holoproteins were detected from PCB-producing E. coli despite that BV should be present as a precursor. BV may be kept at quite low concentration in the PCBproducing E. coli via substrate-channeling from heme oxygenase to PcyA.

### Fluorescence Spectroscopy

Room temperature fluorescence spectra of the thermostable forms of AM1\_C0023g2-PCB and AM1\_C0023g2-BV were measured to evaluate their potentials as fluorescent imaging probes (**Figure 8**). AM1\_C0023g2-BV Pfr form fluoresces with a maximum at 718 nm, whereas AM1\_C0023g2-PCB Pr form fluoresces with a maximum at 679 nm. The fluorescence maximum of AM1\_C0023g2-BV Pfr is red-shifted by 39 nm compared with that of AM1\_C0023g2-PCB Pr, a red shift similar to that found for their absorption spectra maxima. Fluorescence quantum yields of the Pr and Pfr forms were 3.0 and 0.2%, respectively. Fluorescence quantum yield of AM1\_C0023g2-PCB Pr is about two times higher than that of AM1\_1557g2-PCB Pr (Narikawa et al., 2015b).

### Expression of AM1\_1557g2 and AM1\_C0023g2 in Mammalian Cells

We examined the fluorescence of AM1\_1557g2-PCB and AM1\_C0023g2-PCB in live mammalian cells. To precisely compare the fluorescence intensities of the CBCRs, we fused GFP to the CBCRs as an expression control (**Figure 9A**). HeLa cells were transfected with GFP-fused AM1\_1557g2 and

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Anders, K., and Essen, L.-O. (2015). The family of phytochrome-like photoreceptors: diverse, complex and multi-colored, but very useful. Curr. Opin. Struct. Biol. 35, 7–16. doi: 10.1016/j.sbi.2015.07.005

AM1\_C0023g2 and treated with or without 20 µM PCB. Confocal fluorescence images of GFP-fused AM1\_1557g2-PCB and AM1\_C0023g2-PCB show bright green fluorescence of GFP, indicating that both GFP-fused AM1\_1557g2 and AM1\_C0023g2 were successfully expressed in the mammalian cells (**Figure 9B** upper left and right). Almost the same brightness of GFP indicates that the expression levels of GFP-fused AM1\_1557g2 and AM1\_C0023g2 were similar (**Figure 9B** upper left and right). No near-infrared fluorescence was detected from both cells without addition of PCB (**Figure 9C**). Low fluorescence quantum yield and/or low intrinsic BV level may be the reason for this result. On the other hand, the near-infrared fluorescence was detected from both cells after exposure to PCB. The nearinfrared fluorescence of AM1\_C0023g2-PCB was much higher than that of AM1\_1557g2-PCB (**Figure 9B** lower right and left). Relative intensities of AM1\_1557g2-PCB and AM1\_C0023g2- PCB normalized to GFP intensities were 0.31 ± 0.10 and 0.49 ± 0.17, respectively (**Figure 9C**). This is the first report detecting fluorescence from PCB-binding CBCRs expressed in the mammalian cells. Their quantum yields are not so high, but comparable to those of the native phytochromes and CBCRs (Sineshchekov, 1995; Chen et al., 2012; Rockwell et al., 2012c; Pennacchietti et al., 2015). Thus, we can consider that AM1\_C0023g2-PCB is appropriate for platform of fluorescent probe development as a starting material.

### AUTHOR CONTRIBUTIONS

RN, MI, and MS designed the research. KF, TN, YA, TY, N-N-W, and RN performed experiments. N-N-W constructed plasmids. KF and RN performed protein purification and spectroscopic analyses. TN and YA performed fluorescence detection from mammalian cells. KF, TN, MS, and RN wrote the manuscript.

### FUNDING

This work was supported by Precursory Research for Embryonic Science and Technology, Japan Science and Technology Agency, 4-1-8 Honcho Kawaguchi, Saitama 332-0012 (to RN) and Grantsin-Aid for Young Scientists (to RN, 26702036).

### ACKNOWLEDGMENTS

We thank Prof. V.K. Deo (Shizuoka University) for critical reading of the manuscript. We thank Prof. K. Awai for experimental supports and helpful discussion (Shizuoka University).

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Burgie, E. S., Walker, J. M., Phillips, G. N. J., and Vierstra, R. D. (2013). A photolabile thioether linkage to phycoviolobilin provides the foundation for the

blue/green photocycles in DXCF-cyanobacteriochromes. Structure 21, 88–97. doi: 10.1016/j.str.2012.11.001

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The reviewer Y-IlP and handling Editor declared a current collaboration and the handling Editor states that the process nevertheless met the standards of a fair and objective review.

Copyright © 2016 Fushimi, Nakajima, Aono, Yamamoto, Ni-Ni-Win, Ikeuchi, Sato and Narikawa. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# RNA-seq Profiling Reveals Novel Target Genes of LexA in the Cyanobacterium Synechocystis sp. PCC 6803

Ayumi Kizawa1 , Akihito Kawahara2 , Yasushi Takimura2 , Yoshitaka Nishiyama1 and Yukako Hihara1, 3 \*

*1 Department of Biochemistry and Molecular Biology, Graduate School of Science and Engineering, Saitama University, Saitama, Japan, 2 Biological Science Laboratories, KAO Corporation, Wakayama, Japan, 3 Core Research of Evolutional Science and Technology, Japan Science and Technology Agency, Saitama, Japan*

LexA is a well-established transcriptional repressor of SOS genes induced by DNA damage in *Escherichia coli* and other bacterial species. However, LexA in the cyanobacterium *Synechocystis* sp. PCC 6803 has been suggested not to be involved in SOS response. In this study, we performed RNA-seq analysis of the wild-type strain and the *lexA*-disrupted mutant to obtain the comprehensive view of LexA-regulated genes in *Synechocystis*. Disruption of *lexA* positively or negatively affected expression of genes related to various cellular functions such as phototactic motility, accumulation of the major compatible solute glucosylglycerol and subunits of bidirectional hydrogenase, photosystem I, and phycobilisome complexes. We also observed increase in the expression level of genes related to iron and manganese uptake in the mutant at the later stage of cultivation. However, none of the genes related to DNA metabolism were affected by disruption of *lexA*. DNA gel mobility shift assay using the recombinant LexA protein suggested that LexA binds to the upstream region of *pilA7*, *pilA9*, *ggpS,* and *slr1670* to directly regulate their expression, but changes in the expression level of photosystem I genes by disruption of *lexA* is likely a secondary effect.

#### Keywords: cyanobacteria, LexA, RNA-seq, Synechocystis, transcriptome

## INTRODUCTION

The LexA protein in Escherichia coli has been well-characterized as the key regulator of the SOS response induced by DNA damage (Butala et al., 2009). Under non-stress conditions, LexA binds to the promoter regions of more than 40 genes involved in the SOS response and represses their expression. When DNA is damaged, LexA undergoes autoproteolytic cleavage upon association with RecA protein activated through binding of single-stranded DNA fragments. As a consequence of auto-cleavage of the Ala84-Gly85 peptide bond carried out by Ser119 and Lys156, LexA loses DNA binding activity, thereby inducing the SOS response.

Genes encoding LexA homologs are highly conserved in bacterial genomes and LexA-dependent transcriptional regulation of genes involved in DNA repair has been reported in various bacterial species (Erill et al., 2007; Butala et al., 2009), indicating that the regulation of SOS regulon by LexA might be a universal adaptation strategy of bacteria to DNA damage. However, LexA homologs

#### Edited by:

*Takashi Osanai, Meiji University, Japan*

#### Reviewed by:

*Lei Chen, Tianjin University, China Rei Narikawa, Shizuoka University, Japan*

\*Correspondence: *Yukako Hihara<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology*

> Received: *07 January 2016* Accepted: *04 February 2016* Published: *19 February 2016*

#### Citation:

*Kizawa A, Kawahara A, Takimura Y, Nishiyama Y and Hihara Y (2016) RNA-seq Profiling Reveals Novel Target Genes of LexA in the Cyanobacterium Synechocystis sp. PCC 6803. Front. Microbiol. 7:193. doi: 10.3389/fmicb.2016.00193* in several cyanobacterial species were suggested not to be involved in the typical E. coli-type SOS regulation. In Anabaena sp. PCC 7120, auto-cleavage of the Ala84-Gly85 bond of LexA does not occur at physiological pH even in the presence of activated RecA (Kumar et al., 2015). In the case of Synechocystis sp. PCC 6803 (S.6803), LexA lacks the conserved Ala-Gly autocleavage site and the serine of the Ser-Lys dyad required for autocleavage activity (Patterson-Fortin et al., 2006) and auto-cleavage of LexA in S.6803 has not been reported so far. DNA microarray analysis revealed that LexA depletion did not affect the expression level of genes involved in DNA metabolism (Domain et al., 2004).

The cellular processes regulated by LexA in S.6803 have been implied by studies reporting isolation of LexA as a binding factor to the promoter region of specific genes, such as the hoxEFUYH operon encoding bidirectional hydrogenase (Gutekunst et al., 2005; Oliveira and Lindblad, 2005), crhR encoding RNA helicase (Patterson-Fortin et al., 2006), and sbtA encoding sodium-dependent bicarbonate transporter (Lieman-Hurwitz et al., 2009). Domain et al. (2004) performed DNA microarray analysis of the LexA-depleted strain and found that most of genes affected were previously reported to be regulated by the availability of inorganic carbon (Wang et al., 2004). Kamei et al. (2001) reported that the lexA-disrupted mutant of the motile strain of S.6803 (denoted PCC strain) showed non-motile phenotype. DNA microarray analysis revealed that expression of the pilA genes encoding the subunits of the type IV piluslike structure was lowered in the mutant. Although regulation of various cellular processes has been suggested, we currently have still a fragmentary understanding of the function of LexA in S.6803.

DNA microarray analysis has been the most popular methods of genome-wide transcriptome profiling. However, it has been supplanted by RNA-seq analysis in which isolated transcripts are converted into the complementary DNA (cDNA) followed by direct sequence in a massively parallel DNA sequencing-based approach. The advantages of RNA-seq over DNA microarray are its higher resolution and better dynamic range of detecting differential gene expression (Zhao et al., 2014). In order to obtain the comprehensive view of LexA-regulated genes in S.6803, here we performed RNA-seq analysis of the wild-type (WT) strain and the lexA-disrupted mutant. The results of RNA-seq analysis indicate that LexA in S.6803 regulates specific cellular functions such as phototactic motility, accumulation of the major compatible solute glucosylglycerol and subunits of bidirectional hydrogenase, and photosynthetic complexes, but not the SOS response. DNA gel mobility shift assay using the recombinant LexA protein suggested that LexA binds to the upstream region of pilA7, pilA9, ggpS, and slr1670 to directly regulate their expression.

## MATERIALS AND METHODS

### Strains and Culture Conditions

A glucose-tolerant non-motile strain (GT strain) of Synechocystis sp. PCC 6803 was grown at 32◦C in BG-11 medium containing 20 mM HEPES-NaOH, pH 7.0, under continuous illumination at 20µmol photons m−2 s −1 with bubbling of air. The lexA (sll1626)-disrupted mutant (1lexA) was grown under the same conditions, except that 20µg mL−1 kanamycin (Km) was added to the medium. Cell density was estimated by measuring OD730 using a spectrophotometer (model UV-160A, Shimadzu).

### Generation of the lexA (sll1626)-Disrupted Mutant

The coding region of lexA (612 bp, from nucleotide 1319330 to 1318719 according to numbering in CyanoBase) was disrupted by insertion of a kanamycin resistance (Kmr ) cassette. The upstream and downstream fragments including the lexA coding sequence were amplified by PCR from the genomic DNA of the WT strain using the primer sets lexA-F and KmlexA-R (for amplification of 404 bp upstream fragment, from nucleotide 1319525 to 1319122) and Km-lexA-F and lexA-R (for amplification of 394 bp downstream fragment, from nucleotide 1318996 to 1318603; **Table S1**). Kmr cassette was PCR amplified from the pRL161 plasmid using the primer set Km-F and Km-R (**Table S1**). The amplified lexA fragments and Kmr cassette were fused together by the fusion PCR method (Wang et al., 2002) using the primer set lexA-F and lexA-R. The WT strain was transformed with the fusion PCR product and transformants (1lexA mutant) were selected in the presence of Km.

### RNA Gel Blot Analysis

Isolation of total RNA by the hot phenol method and RNA gel blot analyses, using DIG RNA Labeling and Detection Kit (Roche), were performed as described previously (Muramatsu and Hihara, 2003). Template DNA fragments for in vitro transcription to generate RNA probes were prepared by PCR using the primers shown in **Table S1**.

### Immunoblot Analysis

Total proteins were extracted from Synechocystis cells as described previously (Ishii and Hihara, 2008) and separated by 15% (w/v) SDS-PAGE, followed by electroblotting onto PVDF membranes (Immobilon-P; Millipore). Immunodetection was done using a rabbit polyclonal antibody raised against His-LexA recombinant protein. Goat anti-rabbit IgG conjugated to alkaline phosphatase was used as a secondary antibody.

### Determination of Pigment Contents

In vivo absorption spectra of whole cells suspended in BG-11 medium were measured at room temperature using a spectrophotometer (V-650 Spectrometer, JASCO) with ISV-722 integrating sphare. Chlorophyll and phycocyanin contents were calculated from the peak heights of absorption spectra using the equations described in Arnon et al. (1974).

### RNA-seq Analysis

RNA-seq analysis was carried out using cultures at OD730 = 0.5 and OD730 = 1.0 with three biological replicates. WT and 1lexA were inoculated into new media at OD730 = 0.1 and incubated for 50 and 80 h, respectively, to be harvested at OD730 = 0.5. Similarly, WT and 1lexA were inoculated at OD730 = 0.1 and incubated for 70 and 120 h, respectively, to be harvested at OD730 = 1.0. Isolation of total RNA by the hot phenol method was performed as described previously (Muramatsu and Hihara, 2003). To eliminate genomic DNA from total RNA samples, each sample was added with DNase I (TaKaRa) and incubated at 37◦C for 3 h. Total RNA concentration was measured with Nanodrop 2000 (Thermo Fisher Scientific). The Ribo-Zero Magnetic Kit for Bacteria (Epicentre) was used to remove ribosomal RNA from each sample. Concentration and quality of mRNA samples were examined using an Agilent 2100 Bioanalyzer. TruSeq RNA Sample Prep Kit v2 (Illumina) was used for cDNA library construction, and the libraries were sequenced using the Illumina MiSeq system. 12 samples in total were analyzed using two cartridge of MiSeq Reagent Kit v3 (Illumina).

A total of 64 million reads data was obtained from 12 samples. To quantify expression level of each gene, nucleotide sequences of obtained reads were mapped to the genomic sequence of GT-I strain of S.6803 (Kanesaki et al., 2012) (NC\_017038.1; http://www.ncbi.nlm.nih.gov/nuccore/NC\_017038) using CLC Genomics Workbench 7.5.1 software (Qiagen). Raw read counts were divided by length of the transcripts and total number of million mapped reads in each sample to obtain reads per kilobase per million (RPKM) values (Mortazavi et al., 2008). TCC package of R software (Sun et al., 2013) was used to detect the differentially expressed genes between WT and 1lexA. A false discovery rate of <0.01 was considered to be significant.

### Overexpression and Purification of His-lexA

The coding region of the lexA gene was amplified by PCR using the primers lexA-NdeI-F and lexA-XhoI-R (**Table S1**), containing NdeI and XhoI sites at their 5′ end, respectively. The amplified lexA coding fragment was cloned into the pT7Blue Tvector (Novagen), digested with NdeI and XhoI and subcloned into the same restriction sites in pET28a vector (Novagen) to express the LexA protein with an N-terminal 6 × His-tag.

E. coli BL21(DE3) harboring the His-LexA expression construct was grown to an OD600 = 0.6 in 250 mL of 2 × yeast extract-tryptone (YT) medium containing 20µg mL−1 Km at 37◦C and induced with 0.013% of isopropyl β-D-thiogalactoside for 3 h. The cells were pelleted by centrifugation at 5800 g for 2 min, resuspended in 50 mM sodium phosphate buffer, pH 7.4, containing 0.5 M NaCl and 60 mM imidazole, and disrupted by three rounds of sonication with Sonifier 450 (Branson) for 2 min with interval of 1 min on ice. After the removal of whole cells and insoluble material by centrifugation, the soluble protein fraction was filtered through a 0.2µm filter (DISMIC-25CS; ADVANTEC). His-LexA was purified by nickel-affinity column chromatography using a HisTrap FF crude (GE Healthcare). The soluble protein fraction was applied to the column equilibrated with 20 mM phosphate buffer, pH 7.4, containing 0.5 M NaCl and 60 mM imidazole, washed with 20 mM phosphate buffer, pH 7.4, containing 0.5 M NaCl and 80 mM imidazole, and eluted with 20 mM phosphate buffer, pH 7.4, containing 0.5 M NaCl and 300 mM imidazole. Purified His-LexA was desalted by a HiTrap Desalting column (GE Healthcare). Protein composition was examined by 15% (w/v) SDS-PAGE followed by staining with Coomassie Brilliant Blue R-250.

### DNA Gel Mobility Shift Assay

Probes for DNA gel mobility shift assays were obtained by PCR amplification with primers shown in **Table S1** using genomic DNA as a template. The 3′ end of the DNA fragment for each probe was labeled with digoxigenin (DIG)-ddUTP by using the terminal transferase method according to the manufacturer's instructions (DIG gel shift kit 2nd generation; Roche). Gel mobility shift assays were performed by using a DIG gel shift kit 2nd generation (Roche) according to the manufacturer's instruction except that 1 mM DTT was added to the reaction mixture.

## RESULTS

### Characterization of the lexA (sll1626)-Disrupted Mutant

To reveal the function of LexA in GT strain of S.6803, we disrupted the lexA gene by inserting a Kmr cassette within the coding region (**Figure 1A**). Although a fully segregated mutant was not obtained (**Figure 1B**), RNA gel blot and immunoblot analyses revealed that both the lexA transcript (**Figure 1C**) and LexA protein (**Figure 1D**) levels were below the detection limit in the partially segregated mutant (1lexA) grown under normal growth conditions. Under the same conditions, 1lexA displayed several abnormal phenotypes. The doubling time of 1lexA was longer (31.4 h) than that of WT (19.5 h) at log phase, whereas the difference in growth rate between strains became smaller at stationary phase (**Figure 1E**). Amounts of chlorophyll and phycocyanin in 1lexA calculated from the peak heights of cellular absorption spectra were 93 and 80% of WT levels, respectively (**Figure 1F**). Microscopic observation revealed that cell size of 1lexA was heterogeneous and tended to be larger than that of WT (**Figure 1G**).

### RNA-seq Transcriptome Analysis

To investigate the difference in gene expression profile between WT and 1lexA, total RNA was isolated from cultures incubated under normal growth conditions and RNA-seq analysis was performed. **Figure 2** shows MA plots of the gene expression data obtained from cultures at OD730 = 0.5 and OD730 = 1.0. There were 1011 genes differentially expressed between strains at OD730 = 0.5 as shown in magenta (**Figure 2A**). Among them, expression levels of 315 genes were more than two-fold higher and those of 28 genes were more than two-fold lower in 1lexA than in WT (**Table S2**). In the case of WT and 1lexA cells at OD730 = 1.0, there were 447 genes differentially expressed between strains (**Figure 2B**). Among them, expression levels of 360 genes were more than two-fold higher and those of 21 genes were more than two-fold lower in 1lexA than in WT (**Table S3**).

**Table 1** shows the list of genes whose expression was affected by disruption of lexA. The higher resolution and better dynamic range of RNA-seq analysis compared to DNA microarray analysis enabled listing of small ORFs such as ssl1577, ggpR (ssl3076), ssr1251, ssr1473 and ssr3589, and genes with low expression level (low RPKM value) that cannot be detected by previous DNA microarray analyses. Differentially expressed genes can

be categorized into several groups according to related cellular functions as mentioned below.

### Motility-Related Genes

contrast microscopy.

The motile strain of S.6803 exhibits phototactic motility dependent on the type IV-like thick pilus structure (Brahamsha and Bhaya, 2014). In S.6803 genome, there are multiple genes homologous to the pilA gene encoding the subunit of the type IV pilus-like structure. Among them, pilA1 was shown to be responsible for the thick pilus structure, motility, and transformation efficiency (Bhaya et al., 1999; Yoshihara et al., 2001), whereas functions of other pilA-like genes are unknown. We observed that their expression is positively or negatively affected by disruption of the lexA gene. Expression of pilA7-pilA8 was largely enhanced whereas that of pilA9-pilA10-pilA11 and pilA1-pilA2 decreased. The observed decrease in expression level of pilA1 and pilA9-pilA10-pilA11 was consistent with the results of DNA microarray analysis of the 1lexA mutant in the motile PCC strain (Kamei et al., 2001).

Furthermore, we observed that several genes other than pilA involved in motility were affected by disruption of lexA. Expression of pixG-pixH-pixI-pixJ1-pixJ2-pixL (sll0038-0043) encoding regulatory factors involved in positive phototaxis increased in the mutant (**Tables S2**). It has been reported that motility is controlled by cAMP level in S.6803 and inactivation of cya1 encoding adenylate cyclase or sycrp1 encoding cAMP receptor protein results in loss of motility (Terauchi and Ohmori, 1999; Yoshimura et al., 2002a). Although expression of cya1 and sycrp1 itself was not so much affected by disruption of lexA, decrease in expression levels of five genes, pilA9 pilA10-pilA11-slr2018 and cccS (slr1667), out of six genes reported to be decreased by disruption of sycrp1(Yoshimura et al., 2002b) was observed (**Table 1**). cccS is also considered to be related to motility, since its disruption resulted in loss of the thick pili (Yoshimura et al., 2010). We observed expression level of sycrp2 is lower in 1lexA (**Table 1**), although involvement of SYCRP2 in regulation of motility has not been reported.

#### TABLE 1 | Genes with altered expression in the lexA-disrupted mutant.

*(Continued)*

#### TABLE 1 | Continued

*(Continued)*

#### TABLE 1 | Continued

*(Continued)*

#### TABLE 1 | Continued

#### Glucosylglycerol-Related Genes

In S.6803, glucosylglycerol (GG) is a major compatible solute to adapt to high-salt or high-osmotic pressure conditions (Klähn and Hagemann, 2011). A set of genes related to GG biosynthesis (ggp, glp) and uptake (ggt) are organized into several gene clusters such as ggtBCD (slr0529-0531), ggpS-glpD (sll1566-sll1085), ggpP-ggtA (slr0746-0747), and slr1670-glpKspoU-slr1674-hypA1(slr1670-1675) in S.6803 genome (Mikkat and Hagemann, 2000; Klähn et al., 2010). RNA-seq analysis revealed that expression levels of these gene clusters were significantly higher in 1lexA than WT (**Table 1** and **Table S2**). Klähn et al. (2010) reported that a small ORF, ggpR (ssl3076), exists overlapping with the transcription initiation site of ggpS and its promoter region. Expression of ggpR was also induced by disruption of lexA (**Table 1**).

#### Hydogenase-Related Genes

Expression level of the hoxE-hoxF-hoxU-hoxY-hoxH operon encoding subunits of bidirectional NiFe-hydrogenase was lower in 1lexA. This observation is consistent with the previous study reporting that LexA acts as a transcriptional activator for the hox operon (Gutekunst et al., 2005). On the other hand, the expression level of the hypA1 gene involved in hydrogenase maturation increased in 1lexA.

#### Photosynthesis-Related Genes

In S.6803, photosystem (PS) I complex is comprised of 11 subunits and genes encoding these subunits (psa) are dispersed throughout the genome (Kaneko et al., 1996). We found that the expression level of every PSI gene was lower in 1lexA than WT. The expression level of genes encoding subunits of phycobilisome (cpc, apc) was also lower in the mutant, whereas the expression level of genes encoding PSII subunits (psb) was not so much affected by disruption of lexA. Expression levels of chlL-chlN encoding subunits of light-independent protochlorophyllide reductase and that of por encoding lightdependent protochlorophyllide reductase were lower in 1lexA. Both light-dependent and -independent enzymes catalyzing the last step of chlorophyll biosynthesis are likely to be under the control of LexA. On the other hand, expression level of hliA and

hliB encoding high-light inducible proteins was higher in 1lexA than WT.

#### SOS-Response Related Genes

Previous studies suggested that LexA in S.6803 is not involved in the SOS response. Neither lexA nor recA expression was induced upon UV-irradiation (Domain et al., 2004; Patterson-Fortin et al., 2006) and none of DNA metabolism-related genes was listed as genes induced or repressed by LexA depletion (Kamei et al., 2001; Domain et al., 2004). Similarly, induction or repression of DNA metabolism-related genes by disruption of lexA was not observed in our RNA-seq analysis.

### Genes Differentially Expressed in 1lexA at the Later Stage of Growth

Several genes expressed under iron-limiting conditions such as exbB-exbD-fhuA operon involved in inorganic iron uptake (Jiang et al., 2015), futA1 and futA2 encoding subunits of iron transporter (Katoh et al., 2001), and isiA-isiB operon encoding iron-stress inducible proteins (Vinnemeier et al., 1998) were highly induced in 1lexA at OD730 = 1.0 but not in OD730 = 0.5. In the case of mntA and mntC encoding subunits of manganese transporter (Bartsevich and Pakrasi, 1995), their expression level was already higher in 1lexA at OD730 = 0.5 and showed further increase at OD730 = 1.0.

### DNA Gel Mobility Shift Assay