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Until recently, all photosynthetic eukaryotes were considered members of the kingdom Plantae. This is because apart from their ability to capture light energy and fix CO2, they lack many structural and biochemical traits that distinguish plants from protists. Green algae contain the same carotenoids and chlorophyll a and b as land plants, whereas other algae have different accessory pigments and types of chlorophyll molecules in addition to chlorophyll a. Consequently, land plants and closely related green algae are now part of a new monophyletic group called Streptophyta.
The remaining green algae, which belong to a group called Chlorophyta, include more than 7000 different species that live in fresh or brackish water, in seawater, or in snow patches. A few green algae even survive on soil, provided it is covered by a thin film of moisture in which they can live. Periodic dry spells provide a selective advantage to algae that can survive water stress. Some green algae may already be familiar, in particular Spirogyra and desmids. Their cells contain chloroplasts that display a dizzying variety of shapes, and their cell walls contain cellulose, as do land plants. Some green algae are single cells, such as Chlorella and Chlamydomonas, which adds to the ambiguity of green algae classification, because plants are multicellular. Other algae, like Ulva (commonly called sea lettuce), form colonies (Figure 1).
Reproduction of Green Algae
Green algae reproduce both asexually, by fragmentation or dispersal of spores, or sexually, by producing gametes that fuse during fertilization. In a single-celled organism such as Chlamydomonas, there is no mitosis after fertilization. In the multicellular Ulva, a sporophyte grows by mitosis after fertilization (and thus exhibits alternation of generations). Both Chlamydomonas and Ulva produce flagellated gametes.
The charophytes include several different algal orders that have each been suggested to be the closest relatives of the land plants: the Charales, the Zygnematales, and the Coleochaetales. The Charales can be traced back 420 million years. They live in a range of freshwater habitats and vary in size from a few millimeters to a meter in length. The representative genus is Chara (Figure 25.8), often called muskgrass or skunkweed because of its unpleasant smell. Large cells form the thallus: the main stem of the alga. Branches arising from the nodes are made of smaller cells. Male and female reproductive structures are found on the nodes, and the sperm have flagella. Although Chara looks superficially like some land plants, a major difference is that the stem has no supportive tissue. However, the Charales exhibit a number of traits that are significant for adaptation to land life. They produce the compounds lignin and sporopollenin, and form plasmodesmata that connect the cytoplasm of adjacent cells. Although the life cycle of the Charales is haplontic (the main form is haploid, and diploid zygotes are formed but have a brief existence), the egg, and later, the zygote, form in a protected chamber on the haploid parent plant.
The Coleochaetes are branched or disclike multicellular forms. They can produce both sexually and asexually, but the life cycle is basically haplontic. Recent extensive DNA sequence analysis of charophytes indicates that the Zygnematales are more closely related to the embryophytes than the Charales or the Coleochaetales. The Zygnematales include the familiar genus Spirogyra, as well as the desmids. As techniques in DNA analysis improve and new information on comparative genomics arises, the phylogenetic connections between the charophytes and the land plants will continued to be examined to produce a satisfactory solution to the mystery of the origin of land plants.
In bryophytes, gametophytes are the largest and most conspicuous phase of the life cycle.
The gametophyte is haploid and produces haploid gametes by mitosis.
Nutrients are transferred from parent to embryo through placental transfer cells.
Diploid cells called sporocytes undergo meiosis to generate haploid spores.
Female gametangia, called archegonia, produce eggs and are the site of fertilization.
Rhizoids anchor gametophytes to substrate.
Sphagnum, or "peat moss," forms extensive deposits of partially decayed organic material known as peat.
Phloem consists of living cells and distributes sugars, amino acids, and other organic products.
They enable vascular plants to absorb water and nutrients from the soil.
Leaves are categorized by two types:
Microphylls, leaves with a single vein.
Megaphylls, leaves with a highly branched vascular system.
Sterols are vital components of all eukaryotic cells . In higher plants, they play a structural role in cell viability, embryogenesis, pattern formation, cell division, chloroplast biogenesis, and modulation of activity and distribution of membrane-bound proteins such as enzymes and receptors [2, 3]. In addition, sterols are precursors for many signaling molecules that regulate growth and development in plants and animals, such as insect ecdysteroid molting hormones , mammalian steroid hormones , and plant brassinosteroid (BR) hormones .
Sterols belong to a class of isoprenoids derived from isopentenyl pyrophosphate (IPP), a universal precursor of isoprenoids. In animals and fungi, the cytoplasmic mevalonic acid (MVA) pathway is the only route for biosynthesis of IPP, the building block for lanosterol, which is then metabolized into cholesterol in animals and ergosterol in fungi . In higher plants, IPP can be derived via either the MVA pathway or the plastidial 1-deoxyxylulose 5-phosphate or methylerythritol phosphate (MEP) pathway, despite the former being the main contributor to sterol biosynthesis [8, 9]. In Arabidopsis, sterol biosynthetic mutants can be classified into two distinct groups: BR-independent mutants, which are defective in genes in the pathway from cycloartenol to 24-methylenelophenol , and BR-dependent mutants, which are defective in genes at the latter part of the sterol biosynthetic pathway (from 24-methylenelophenol to campesterol, which is considered as the precursor for BRs) . Besides serving as a BR precursor, sterols appear to play distinct signaling functions during plant development, since phenotypes of sterol biosynthetic mutants cannot be rescued by addition of exogenous BR .
Co-regulation of sterol synthesis and fatty acid (FA) production is essential for maintaining the biosynthesis-versus-turnover balance of membranes during cellular growth . In animals, sterols and FAs are coordinately regulated by a feedback system mediated by a conserved family of transcription factors called sterol regulatory element binding proteins (SREBPs), which controls a cascade of biosynthetic enzymes for endogenous cholesterol, FA, triacylglycerol (TAG), and phospholipid [14, 15]. Activated SREBP binds to sterol response elements in the promoter and/or enhancer regions of target genes and induces transcription of at least 30 cholesterol- and lipid-synthesis genes (particularly those encoding rate-limiting enzymes, such as hydroxy-methyl-glutaryl-CoA reductase, HMGR and type I FA synthase, FAS) . Manipulation of this regulatory cascade in transgenic mice resulted in 6- and 22-fold increases in cholesterol content and TAG content, respectively, and consequentially massive fatty livers .
Microalgae are promising feedstock for sustainable and scalable production of biofuel  however, few microalgal strains found in nature are endowed with the wide array of traits demanded by a large-scale and economically competitive production scheme . It is therefore urgent to identify molecules and mechanisms that regulate microalgal growth, development, and stress responses for strain improvement. Sterols in microalgae display enormous diversity due to the high degree of phylogenetic heterogeneity, the vast number of genera, and the long evolutionary distance among many of them [20, 21]. It has long been shown that in microalgae sterol composition varies upon changes in growth stage, light spectrum, or temperature, suggesting important yet largely unknown roles of sterols . Therefore, manipulation of sterol biosynthesis and regulation offers potential for engineering lipid production. Exploration of the sterol-dependent lipogenesis regulatory mechanism in microalgae might provide novel strategies and targets for enhanced lipid production in microalgae. However, little is known about the roles, biosynthesis, and regulation of sterols in microalgae and in particular, whether and how sterol and FA metabolism is co-regulated.
Nannochloropsis spp. are a genus of unicellular photosynthetic microalgae belonging to the heterokonts. They are distributed widely in the marine environment as well as in fresh and brackish waters. These algae are of industrial interest because they grow rapidly and can synthesize large amounts of TAG and high-value polyunsaturated FA (for example, eicosapentaenoic acid) . The genomes of multiple species of oleaginous Nannochloropsis spp. have been sequenced and annotated [23–27]. Employing an oleaginous industrial microalga N. oceanica IMET1 as a model, this study has aimed to determine the sterol composition and biosynthetic pathway in microalgae, to investigate the role of sterol biosynthesis in photosynthesis and growth, to study the influence of light and nitrogen supply, and to probe the effects of sterol levels on FA accumulation. Our findings expand the understanding of sterol function in microalgae and should assist rational genetic or process engineering for microalgae-based production of biofuels or other value-added bioproducts.
Structure and mechanism of the evolutionarily unique plant enzyme chalcone isomerase
Chalcone isomerase (CHI) catalyzes the intramolecular cyclization of chalcone synthesized by chalcone synthase (CHS) into (2S)-naringenin, an essential compound in the biosynthesis of anthocyanin pigments, inducers of Rhizobium nodulation genes, and antimicrobial phytoalexins. The 1.85 Å resolution crystal structure of alfalfa CHI in complex with (2S)-naringenin reveals a novel open-faced β-sandwich fold. Currently, proteins with homologous primary sequences are found only in higher plants. The topology of the active site cleft defines the stereochemistry of the cyclization reaction. The structure and mutational analysis suggest a mechanism in which shape complementarity of the binding cleft locks the substrate into a constrained conformation that allows the reaction to proceed with a second-order rate constant approaching the diffusion controlled limit. This structure raises questions about the evolutionary history of this structurally unique plant enzyme.
2 MATERIALS AND METHODS
2.1 Algal culture conditions
P. tricornutum (CCMP2561), obtained from the culture collection of the Provasoli-Guillard National Center for Culture of Marine Phytoplankton, Bigelow Laboratory for Ocean Sciences, was maintained and cultured in F/2 medium containing 20 g/L sea salt. The strain was denoted as Pt1 with a fusiform morphotype (De Martino et al., 2007 ). Briefly, 10 ml of liquid F/2 medium was inoculated with cells from agar plates and the alga was grown aerobically in 50 ml flasks at 23°C for 6 days (hand shaking twice per day) illuminated with continuous light of 30 µE m −2 s −1 (cool-white fluorescent tube light, from the top). The algal cells were then inoculated at 10% (v/v) into 250 ml flasks for growth with orbital shaking at 150 rpm in an orbital shaker (Zhichu) and constant illumination of 30 µE m −2 s −1 . Algal cells grown to late exponential phase were used as seeds for experiments.
For the treatment of nitrogen deprivation (ND), the seeds were harvested, washed twice with F/2 medium lacking nitrogen, and re-suspended in this medium. Cells re-suspended in F/2 medium (nitrogen replete, NR) were used as the control. Both had a starting cell number of 1.5 × 10 6 ml −1 and were grown for 4 days with orbital shaking at 150 rpm and constant illumination of 30 µE m −2 s −1 . For the comparison between the wild type (WT) and transgenic strains of P. tricornutum, cells were inoculated to a starting cell number of 5 × 10 5 ml −1 and allowed to grow for 12 days with orbital shaking at 150 rpm and constant illumination of 30 µE m −2 s −1 .
2.2 Cloning and in silico analysis of P. tricornutum DGATs
To obtain the full-length coding sequence of PtDGAT1, its untranslated regions were first determined by 5ʹ and 3ʹ rapid amplification of cDNA ends (RACE)-PCR using the SMARTer RACE 5ʹ/3ʹ Kit (TaKaRa). Then, primer pairs were used to amplify the full-length coding sequence, which was verified by sequencing and deposited into NCBI GenBank (accession number MN061782). The sequences of PtDGAT2A through PtDGAT2D and PtDGAT3 were retrieved from NCBI GenBank (JX469835, JQ837823, JX469836, JX469837, and XM_002184438).
Sequence alignment of DGAT polypeptides from various organisms was conducted using ClustalX2.1 (http://www.clustal.org/clustal2/) and the phylogenetic tree was generated using MEGA6 (Tamura et al., 2013 ). Transmembrane helices of PtDGATs were predicted using TMHMM 2.0 (http://www.cbs.dtu.dk/services/TMHMM/).
2.3 RNA isolation and quantitative real-time PCR
Total RNA extraction (from around 10 7 algal cells) and removal of contaminated DNA were conducted by using the plant RNA extraction kit (TaKaRa). After the synthesis of cDNA, quantitative real-time PCR (qPCR) was conducted on a 7500 Fast Real-Time PCR System (Applied Biosystems) with SYBR ® Premix Ex Taq™ II (TaKaRa), following our previously described procedures (Wei et al., 2017 ). Primer sequences used for qPCR are listed in Table S1. The mRNA expression level of PtDGAT genes was normalized using the actin gene as the internal control.
2.4 Functional complementation of PtDGATs in the TAG-deficient yeast
The six PtDGAT genes were PCR amplified using cDNA as template and cloned into the yeast expression vector pYES2-CT (Invitrogen), using the In Fusion Advantage PCR Cloning Kit (TaKaRa). PCR primers for the cloning are listed in Table S1. The recombinant plasmids, once confirmed by sequencing, were each introduced into the TAG-deficient quadruple mutant strain H1246 of Saccharomyces cerevisiae (Sandager et al., 2002 ). The empty vector (EV) pYES2-CT was used as the negative control. The yeast transformants, after verified by Colony PCR, were cultured in SD/-Ura medium containing 2% (w/v) galactose to induce transgene expression. For the free fatty acid experiment, linoleic acid (C18:2), α-linolenic acid (C18:3n3), and eicosapentaenoic acid (C20:5n3) were each fed to yeast cultures upon galactose induction at a concentration of 100 µM, as described by Mao et al. ( 2019 ). After 2 days of cultivation, the yeast cultures were harvested for lipid analysis and fluorescent staining.
2.5 In vitro assay of PtDGATs
The induction of heterologous expression of PtDGAT genes in H1246 and the microsome preparation from these yeast strains containing PtDGATs were conducted according to our previously described procedures (Liu et al., 2017 ). The prepared microsomal fractions were each re-suspended in the storage buffer (50 mM Tris-HCl, pH 7.5, 10% glycerol) for subsequent in vitro assay.
The in vitro DGAT assay was performed in a 200 µl reaction system, which consists of 20 µg microsomal protein, 250 µM of both substrates acyl-CoA and DAG, and 10 mM MgCl2 in potassium phosphate buffer (Liu et al., 2017 ). DAG (16:1/16:1) was used as the acyl acceptor, while 16:0-CoA, 16:1-CoA, and C20:5-CoA were used as the acyl donor all were purchased from Larodan Fine Chemicals.
2.6 Overexpression of PtDGATs in P. tricornutum
The coding sequences of PtDGAT1, PtDGAT2A through PtDGAT2D and PtDGAT3 were each subcloned into the P. tricornutum expression vector peGFP (Zhang & Hu, 2014 ), using the In Fusion Advantage PCR Cloning Kit (TaKaRa). PCR primers for the cloning are listed in Table S1. The transformation of P. tricornutum via electroporation followed our previously described protocols (Zhang & Hu, 2014 ). Putative transformants selected on F/2 solid medium containing 75 μg/ml zeocin were verified by genomic PCR. Then, qPCR was performed to determine the expression level of transgenes in the selected transformants. Two strains with the highest expression levels for each transgene were chosen for further experiments.
2.7 Fluorescent and confocal microscopy analyses
For the observation of neutral lipids in P. tricornutum and yeast cells, they were first stained with the fluorescence BODIPY™ 505/515 (Invitrogen) with a 1 µg/ml working concentration for 10 min at room temperature, and then visualized under an Olympus BX51 Fluorescence Microscope (Olympus) with excitation at 488 nm and emission between 505 and 530 nm. For subcellular localization of PtDGATs, the corresponding transgenic algal strains were visualized under a Leica TCS SP8 laser scanning confocal microscope. GFP fluorescence was observed with excitation at 488 nm and emission at 500–525 nm, and chlorophyll autofluorescence was observed with excitation at 488 nm and emission at 650–750 nm. To observe the nucleus of algal cells, they were stained with Hoechst 33342 (Invitrogen) at a concentration of 5 µM for 30 min at room temperature and then visualized with excitation at 405 nm and emission at 425–475 nm.
2.8 Analytical methods
Cell counting using a hemocytometer under a light microscope and gravimetrical determination of dry weight for P. tricornutum followed our previously described procedures (Liu et al., 2019 ). The maximum quantum yield of PSII, Fv/Fm or Fm − Fo/Fm, was measured on a pulse amplitude-modulated (PAM) fluorometry (Water-PAM, Walz), using 2 hr dark-adapted algal cell (Liu et al., 2019 ): Fo was recorded under a weak light (<10 μmol m −2 s −1 , peaking at 650 nm) and Fm was under a saturating pulse (0.8 s) of red light (3,000 μmol m −2 s −1 , peaking at 660 nm). Protein content in P. tricornutum cells was determined according to Meijer and Wijffels ( 1998 ), using lyophilized algal samples. Carbohydrate content in P. tricornutum cells was determined by the colorimetric method (Renaud et al., 1999 ), after hydrolysis of the lyophilized algal samples with 4 M H2SO4.
Lipids from yeast and P. tricornutum cells were extracted with chloroform–methanol (2:1, v/v) as previously described by Mao et al. ( 2019 ). For thin-layer chromatography (TLC) analysis, the extracted lipids were separated on a Silica gel 60 TLC plate (Merck) using a mixture of hexane/tert-butylmethyl ether/acetic acid (80/20/2, by vol) as the mobile phase (Liu et al., 2016 ). After separation, TAG on the TLC plate was visualized with iodine vapor, recovered, and transesterified with 1.5% sulfuric acid in methanol at 85°C for 2.5 hr. Total lipids from P. tricornutum and yeast cultures and the reaction products from in vitro DGAT assays were directly transesterified.
Fatty acid methyl esters prepared from the transesterification of lipids were analyzed by using an Agilent 7890 capillary gas chromatograph equipped with a 5975 C mass spectrometry detector and a HP-88 capillary column (60 m × 0.25 mm Agilent Technologies), following our previously described procedures (Liu et al., 2016 ).
Plant material and growth conditions
Mutant and transgenic plants of Arabidopsis (Arabidopsis thaliana) were in the Col-0 wild-type background. Arabidopsis seeds were sown on compost, stratified for 3 days at 4 °C and grown at 20 °C, under ambient CO2 (ca. 400 μmol/mol), at 70% relative humidity and under 100 μmol photons/m 2 /s in 12-h light/12-h dark cycles, unless otherwise stated.
For analyses of transgenic lines, homozygous insertion or wild-type out-segregant lines (T3) were compared. Where wild-type out-segregants were not available, homozygous insertion lines were compared with Col-0 plants from seed stocks of the same age generated under similar conditions. Tobacco plants (Nicotiana benthamiana L.) were cultivated under glass house conditions (minimum 20 °C, natural light supplemented to give at least 12-h light). Venus-tagged proteins were expressed in wild-type Chlamydomonas strain cMJ030 (CC-4533)(Zhang et al., 2014 ). Cells were maintained in constant low light (
10 μmol photons/m 2 /s) at RT on 1.5% (w/v) agar plates containing Tris–acetate–phosphate (TAP) (Kropat et al., 2011 ). For imaging, cells were grown in liquid TAP media to a concentration of 10 6 cells/mL, pelleted by centrifugation (1000 g, 4 min), resuspended in Tris–phosphate (T-P) minimal media (Kropat et al., 2011 ) and grown for 24 h in ambient CO2 before imaging.
Cloning and expression of CCM components in Chlamydomonas
The open reading frames (ORFs) of Chlamydomonas genes were expressed in frame with Venus from the PsaD promoter using the pLM005 vector. ORFs were amplified from genomic DNA using Phusion Hotstart II polymerase (Thermo Fisher Scientific, www.thermofisher.com) with the respective oligos in Table S1. HpaI-cut pLM005 vector and PCR products were gel purified and assembled by Gibson assembly (Gibson et al., 2009 ). Due to the large gene length of HLA3, it was cloned in two fragments then assembled in the pLM005 vector by Gibson assembly. The pLM005 vector contains the AphVIII gene for paromomycin resistance in Chlamydomonas and ampicillin resistance for bacterial selection. All construct junctions were verified by Sanger sequencing. Constructs were transformed into Chlamydomonas by electroporation as in Zhang et al. ( 2014 ). Briefly, 250 μL of 2 × 10 8 cells/mL was transformed with 14.5 ng/kbp of EcoRV-cut plasmid at 16 °C. Cells were spread on 86 mL TAP agar plates containing paromomycin (20 μg/mL) and kept in low light (
10 μmol photons/m 2 /s) until colonies were
2–3 mm in diameter. Plates were screened for fluorescent colonies using a Typhoon TRIO fluorescence scanner (GE Healthcare, www.gelifesciences.com) with excitation/emission wavelengths 532 nm/520–555 nm for Venus and 633 nm/630–670 nm for chlorophyll autofluorescence.
Cloning and expression of CCM components in tobacco and Arabidopsis
Genes were cloned from cDNA derived from Chlamydomonas (strain CC-4886, Chlamydomonas Resource Center). Primers were designed from sequences available on Phytozome v10.2 (Chlamydomonas reinhardtii v5.5 [Augustus u11.6], http://phytozome.jgi.doe.gov/pz/portal.html#!info?alias=Org_Creinhardtii) (see Table S1 for oligo details). Gene sequences for LCIA and HLA3 were codon-optimized for expression in higher plants and synthesized de novo (DNA2.0, CA, USA) (Figure S7), then cloned into Gateway entry vectors (pCR ® 8/GW/TOPO ® TA Cloning ® Kit) using Platinum ® Taq DNA Polymerase High Fidelity according to the manufacturer's instructions (Invitrogen ™ Life Technologies, www.lifetechnologies.com) and subsequently cloned into the destination binary vectors pK7FWG2,0 (Karimi et al., 2002 ) or pGWB5 (Nakagawa et al., 2009 ). Gibson assembly was used to generate transit peptide gene fusions. Binary vectors were transformed into Agrobacterium tumefaciens (AGL1) for transient gene expression in tobacco leaves (Schöb et al., 1997 ) or stable insertion in Arabidopsis plants by floral dipping (Clough and Bent, 1998 ). Co-expression studies were performed with the WAVE131 vector of the ‘wave’ marker set (Geldner et al., 2009 ) and the mt-rb vector (Nelson et al., 2007 ) for plasma membrane and mitochondria localization, respectively.
DNA extraction, PCR and protein analysis
For screening transgenic Arabidopsis lines, genomic DNA was extracted from mature, nonflowering rosettes of T1 plants as described in Li and Chory (Li and Chory, 1998 ). PCRs were performed as in McCormick and Kruger ( 2015 ). Where possible, the location of gene inserts was confirmed by TAIL PCR as described by Liu et al. ( 1995 ). Homozygous insertion lines were identified in the T2 generation either by PCR or by seedling segregation ratios on kanamycin-containing Murashige and Skoog (MS) medium (0.5x) plates.
Relative levels of LCIA: GFP and HLA3: GFP proteins in leaves were confirmed by immunoblot. Approximately 10 μg protein from whole 28-d-old rosettes (100 mg fresh weight) was fractionated by SDS-PAGE on a 10% (w/v) acrylamide: bisacrylamide (40:1) gel transferred to PVDF membrane, probed with mouse anti-GFP IgG2a at 1:1,000 dilution (Santa Cruz, http://www.scbt.com/) and visualized using an HRP-conjugated goat anti-mouse IgG2a at 1:50 000 dilution. HRP activity was detected using Supersignal Ultra (Pierce, www.piercenet.com) according to the manufacturer's instructions.
Oocyte expression and bicarbonate uptake assays
Synthesized gene sequences for mature LCIA (mLCIA) or HLA3 lacking a stop codon were cloned into the expression vector Vivid Colors ™ pcDNA ™ 6.2/EmGFP (Invitrogen ™ Life Technologies) to generate mLCIA- or HLA3-GFP fusions. For LCIA, the N-terminal transit sequence peptide (73 aa) was removed and replaced with the sequence ‘GACATG’ to add a Kozak sequence and new start codon. For HLA3, ‘GAC’ was added immediately upstream of the start codon. To generate equivalent vectors lacking a fluorescent tag, Gibson assembly was used to add a stop codon to mLCIA or HLA3 and remove the GFP sequence (720 bp). Plasmids were linearized by AvrII or StuI (Roche, www.roche.co.uk) and capped mRNA was synthesized using the mMESSAGE mMACHINE ® T7 Transcription Kit (Ambion ® Life Technologies) according to the manufacturer's instructions. Mature LCIA or HLA3 mRNA was expressed in oocytes as described by Feng et al. ( 2013 ). Xenopus oocytes were injected with 50 nL of mRNA (1 μg/μL) or diethyl pyrocarbonate (DEPC)-treated water as a control. Bicarbonate uptake assays and confocal imaging were performed 3 d after injection in a protocol adapted from Mariscal et al. ( 2006 ). Oocytes were incubated in fresh MBS media (88 m m NaCl, 1 m m KCl, 2.4 m m NaHCO3, 0.71 m m CaCl2, 0.82 m m MgSO4 and 15 m m HEPES, pH 7.4), containing 0.12 m m NaHCO3 (1.85 GBq/mol NaH 14 CO3). After 10 min, the oocytes were washed three times with ice-cold MBS and lysed in 200 μL SDS (10% [w/v]), and the radioactivity retained in individual oocytes was measured.
Chlorophyll was extracted from powdered leaf discs in ice-cold 80% (v/v) acetone and 10 m m Tris–HCl, and concentration was measured according to Porra et al. ( 1989 ).
Measurement of photosynthetic parameters
Gas exchange rates were determined using a LI-6400 portable infrared gas analyser (LI-COR Biosciences, http://www.licor.com/) on either the sixth or seventh leaf of 35- to 45–d-old mature, nonflowering rosettes grown in large pots to generate leaf area sufficient for gas exchange measurements (Flexas et al., 2007 ). The response of net photosynthetic CO2 assimilation (A) to substomatal CO2 concentration (Ci) was measured by varying the external CO2 concentration from 0 to 1000 μmol/mol under a constant photosynthetic active radiation of 1500 μmol photons/m 2 /s (provided by a red–blue light source attached to the leaf chamber). Gas exchange data were corrected for CO2 diffusion as in Bellasio et al. ( 2015 ). Leaf temperature and chamber relative humidity were maintained at 21 °C and 70%, respectively. To calculate maximum carboxylation rate (Vc,max), maximum electron transport flow (Jmax) and mesophyll conductance (gm), the A/Ci data were fitted to the C3 photosynthesis model (Farquhar et al., 1980 ) with modifications to include estimations for gm as described by Ethier and Livingston ( 2004 ).
Confocal laser scanning microscopy
A Leica TCS SP2 laser scanning confocal microscope (Leica Microsystems) with a water immersion objective lens (HCX IRAPO 25.0x0.95) was used for imaging leaves and oocytes. Excitation/emission wavelengths were 488 nm/500–530 nm for GFP, 543 nm/590–620 nm for mCherry and 488 nm/680–750 nm for chlorophyll autofluorescence. Images were acquired using Leica LAS AF software (http://www.leica-microsystems.com/). Prior to imaging Venus-tagged proteins in Chlamydomonas, 15 μL of cells were added to a well of a 96-well optical plate (Brooks Life Science Systems, http://www.brooks.com) and covered with 150 μL of 1.5% low melting point agarose containing T-P (
35 °C). For mitochondria staining, CCP1: Venus- and CCP2: Venus-expressing lines were grown in liquid T-P media with paromomycin (2 μg/mL) to a concentration of 2–4 × 10 6 cells/mL. Cultures were incubated with MitoTracker Red CMXRos (Thermo Fisher Scientific) to a final concentration of 1 μ m for 10 min, then spotted on a polylysine-coated slide for imaging. Cells were imaged using a custom adapted confocal microscope (Leica DMI6000) with settings at 514 nm/532–555 nm for Venus, 561 nm/573–637 nm for staining by Mitotracker Red CMXRos and 561 nm/665–705 nm for chlorophyll autofluorescence. Images were analysed using Fiji software (http://fiji.sc/Fiji).
Variations in response between genotypes were assessed by either analysis of variance (ANOVA) or Student's t-tests followed by Tukey's honest significant difference (HSD) post hoc test (SPSS Statistics 18, http://www.ibm.com/). Differences for which P < 0.05 are considered significant.
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Here, we identified 92 COMT genes from blueberry and 425 COMT genes from 15 other species. According to phylogenetic analysis of COMTs, we divided the COMTs into two groups, which indicated the existence of two ancestor genes. DSD and WGD were revealed to be the major forces of blueberry evolution. The Ka/Ks ratios of the gene duplication patterns for the COMTs from the four plant species were less than 1, indicating that the COMTs have experienced strong purifying selection. According to the qRT-PCR results for 22 VcCOMTs, VcCOMT40, VcCOMT92 were highly expressed and may play important roles in the synthesis of lignin of blueberry fruit. The results of this study will build foundations for breeding blueberry cultivars with higher fruit firmness and longer shelf life.
We acknowledge funding of this research project by the Research Council of Norway (RCN) and the University of Hamburg (Hamburg, Germany). We are grateful to Prof. C. Benning (Michigan State University, East Lansing, United States) for providing the expression vector pNoc ox Venus. We also would like to thank Elke Wölken (Department of Aquatic Ecophysiology and Phycology, University of Hamburg) for analyses of immunogold-labeled N. oceanica transformants by transmission electron microscopy.
aa, amino acid ALNS, allantoin synthase ASW, artificial sea water At, Arabidopsis thaliana CaMV, cauliflower mosaic virus DC, decarboxylase DECR, 2,4-dienoyl-CoA reductase DHNS, 1,4-dihydroxy-2-naphthoyl-CoA synthase dpt, days post transformation EMB8, embryogenesis-associated protein 8 EPA, eicosapentaenoic acid EYFP/GFP, enhanced yellow/green fluorescent protein HIT, histidine triad family protein HIUase, 5-hydroxyisourate hydrolase IndA, indigoidine synthase A MDH, malate dehydrogenase MLS, malate synthase Ng, Nannochloropsis gaditana OHCU, 2-oxo-4-hydroxy-4-carboxy-5-ureidoimidazoline PEX, peroxin PfkB, 6-phosphofructokinase PGL3, 6-phosphogluconolactonase 3 PKT, peroxisomal 3-ketoacyl thiolase PTS1/2, peroxisomal targeting signal type 1/2 PUFA, polyunsaturated fatty acid PUKI, pseudouridine kinase PUMY, pseudouridine monophosphate glycosylase TEM, transmission electron microscopy.
Proteome profile changes in P. patens under ABA treatment
Changes in the proteome profile between control and ABA treated plants suggested that exogenous ABA could trigger one or more responses in P. patens. In order to ensure statistically results, the experiments were carried out in triplicate. After CBB R-250 staining, more than 1,300 protein spots could be detected for each sample. Representative gels from control and ABA-treated plants and proteins showing altered abundance are presented in Figure Figure1. 1 . Two-dimensional images were analyzed using ImageMaster™ 2D Platinum software. The volume percentage of each spot was estimated and compared across the gels. In the samples following ABA treatment, we repeatedly analyzed 89 protein spots, whose relative abundance was at least 1.5-fold different from the control. The proteins associated with 65 of these protein spots were subsequently identified using LC-MS/MS. Among them, 13 protein spots (D1-D13) were down-regulated, 52 protein spots (U1-U52) were up-regulated by ABA treatment including four protein spots (U22, U24, U40, U43) which were only present in the ABA-treated samples (Figure (Figure1). 1 ). Quantitative analysis indicated that the abundance of these proteins changed [see Additional File 1: Supplemental Table S1] in response to ABA treatment (Figure (Figure2) 2 ) suggesting that different physiological and biochemical processes are modified following ABA treatment.