Transcriptomic Analysis Reveals the Pathways Associated with

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Transcriptomic analysis reveals the pathways associated with resisting and degrading microcystin in Ochromonas Lu Zhang, Kai Lyu, Na Wang, Lei Gu, Yunfei Sun, Xuexia Zhu, Jun Wang, Yuan Huang, and Zhou Yang Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b03106 • Publication Date (Web): 04 Sep 2018 Downloaded from http://pubs.acs.org on September 4, 2018

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Environmental Science & Technology

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Transcriptomic analysis reveals the pathways associated with resisting and degrading

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microcystin in Ochromonas

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Lu Zhang†, Kai Lyu†, Na Wang†, Lei Gu†, Yunfei Sun†, Xuexia Zhu†, Jun Wang†, Yuan

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Huang†, Zhou Yang*,†, ‡

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†

Jiangsu Province Key Laboratory for Biodiversity and Biotechnology, School of

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Biological Sciences, Nanjing Normal University, 1 Wenyuan Road, Nanjing 210023,

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China

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‡

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Guangzhou 510632, China

Department of Ecology, College of Life Science and Technology, Jinan University,

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* Corresponding author: Zhou Yang

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TEL: +86-25-85891671.

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E-mail: [email protected]

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ACS Paragon Plus Environment


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Abstract

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Toxic Microcystis bloom is a tough environment problem worldwide. Microcystin is

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known as highly toxic and easily-accumulated secondary metabolites of toxic

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Microcystis and threaten water safety. Biodegradation of microcystin by protozoan

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grazing is a promising and efficient biological method, but the mechanism in this

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process is still unclear. The present study aimed to identify potential pathways

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involved in resisting and degrading microcystin in flagellates through transcriptomic

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analyses. A total of 999 unigenes were significantly differentially expressed between

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treatments with flagellates Ochromonas fed on microcystin-producing Microcystis

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and microcystin-free Microcystis. These dysregulated genes were strongly associated

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with translation, carbohydrate metabolism, phagosome, and energy metabolism.

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Upregulated genes encoding peroxiredoxin, serine/threonine-protein phosphatase,

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glutathione S-transferase, HSP70, and O-GlcNAc transferase were involved in

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resisting microcystin. In addition, genes encoding cathepsin and GST and genes

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related to inducing ROS were all upregulated, which highly probably linked with

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degrading microcystin in flagellates. The results of this study provided a better

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understanding of transcriptomic responses of flagellates to toxic Microcystis as well

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as highlighted a potential mechanism of biodegrading microcystin by flagellate

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Ochromonas, which served as a strong theoretical support for control of toxic

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microalgae by protozoans.

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Key words: Degradation pathway; Microcystin; Ochromonas; RNA-seq; Microcystis

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microcystin-producing Microcystis TM

Ochromonas Treatment Time

Resistant mechanism

Control

RNA-Seq Perc ent of ge nes

Microcystis dynamics (MC concentration)

bi ologic al _proc ess

cell cell j nuc ti on e tx c ra cellu ellpa tr l ar ext ar cel re gi on macro lul arer gi nop m lo a ecul tr arc o m pl ex memb memb ra ne memb ar ne ar ne pa tr -e cn los ed l um en or ga ne lle or ga ne l le supar pa tr mol ec lu arﬁ be r sym pl ast vi ri o n vi ir o ant p a n i oxi tr ad nt a ctiv i ty catal bi ndi ng ty ic el ectro a ctiv nc i ty arri er met alloc a ctiv i ty mol ah pe rone ecul ncu arfunc l eic a ctiv mol i ty a ci d t ionre ecul bi n art id gnt gul ar ns at ro ra ns duc era ctiv cri pt ionfa i ty unt ri ct roa ent ctivi re se ty rvoi signa ra ctiv lt ra ns du i ty struc c er tura a ctiv l mol i ty ecul e a ctiv tra ns i ty port era ctiv i ty

bi ol oig cala pone bi ol d nt or ogi cal he sion ag ni re gul zatio at ion norbi goe ne cellu si s l arpr oc ess de to de ve xi ﬁcati lopm no ent al proc ess i mmu grow ne sy th s t em porc ess loc metab alizati o n ol i mul cporc mul t i-or en ga ga ni ess t icell ul ar t ive sm pr re g or g oc l a u pos it ni smal ess ation ive ofbi porc re glu lo ogi ess ation cal ofbi porc re gul ess lo ogi ationo cal p fbi orc lo ogi ess cal porc ess er prodcu re produ ti on c t ive porc er sp ons ess et os timu hryt l us hm i cproc ess singl si ng e-or a l in ga ni g sm porc ess

Experimental design

cel lu l arc om

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Microcystis (microcystins)

Page 3 of 48 Environmental Science & Technology

Abstract Art Differentially expressed genes 100

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577

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c el lul ar_compone nt m ol ec ula r_func t ion

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N -vs -T

U p-R e D ow

fed on

Degradation process

fed on

microcystin-free Microcystis NM

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1. Introduction Influences of climate change and water eutrophication on aquatic ecosystem have

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been closely focused on worldwide, due to increasing appearance of cyanobacterial

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blooms.1,2 Extreme proliferation of cyanobacteria has become a severe problem in

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freshwater, where massive cyanobacteria disturb the ecosystem function and plankton

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assemblages and the release of accompanied metabolites increases the risk to water

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safety.3,4 Toxic Microcystis, one of the most prevalent bloom-forming cyanobacteria,

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can produce microcystin (MC), a major type of toxic secondary metabolites. The toxic

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mechanisms of microcystin mainly include inhibition of protein phosphatases (PP1

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and PP2A) in eukaryotic organisms,5 disruption of cytoskeleton,6 and induced

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oxidative stress (Amado and Monserrat, 2010).7 In several drinking water sources,

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toxic Microcystis blooms frequently occur,8,9 and microcystin concentration

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significantly exceeds the maximum allowable value of 1 µg L-1 for drinking water, as

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proposed by the World Health Organization, thereby posing a serious hazard to human

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and animal health. Hepatic illness and endocrine-disrupting effects on reproduction

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attribute to microcystin.10,11

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To date, various methods have been developed to reduce abundance of toxic

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Microcystis and remove microcystin. For instance, potassium permanganate and

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hydrogen peroxide can cause cell rupture to reduce cyanobacterial biomass;12,13

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ferrate and hydroxyl radical mainly oxidize aromatic ring, diene in Adda, and the

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double bond of the methyldehydroalanine to degrade microcystin.14,15 In addition to

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these chemical and physical methods, bacteria and aquatic herbivorous predators are

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generally used in removal of cyanotoxins and control of cyanobacterial populations

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among biological methods.16 Especially, based on the fact that predation plays a

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critical and efficient role in transferring the matter and energy of preys to high trophic

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levels, herbivorous zooplanktons and fishes can ingest cyanobacteria to some extent,

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however, these large-sized grazers were usually threatened by cyanotoxins,17-19 due to

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lack of strong resistance and degradation capability. Previous studies have suggested

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that the MC were accumulated in liver and muscle of fish through trophic transfer of

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MC from zooplankton,20,21 which would further threaten the health of final consumers.

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Thus, these large-sized predators are not qualified to reduce toxic cyanobacterial

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biomass and degrade toxins.

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In contrast, some species of protozoans, a type of small-sized zooplankton, not

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only have inhibition effects on Microcystis populations by grazing but also mostly

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show strong resistance to MC. Within these protozoans (Supporting Information, see

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Table S1), amoebae (e.g., Acanthamoeba castellanii and naked amoeba) can graze

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unicellular or colonial Microcystis and resist MC without degradation capability;22,23

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ciliates (e.g., Nassula sp.) can decrease MC concentration but cannot degrade Adda

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group of MC;24 flagellates (e.g., Ochromonas sp., Poterioochromonas sp., Diphylleia

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rotans, and Monas sp.) can ingest Microcystis, accompanied with degrading MC.25-28

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Moreover, flagellates grazing Microcystis facilitates chlorophytes dominance, thereby

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improving phytoplankton community and water quality.29 Consequently, flagellates

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play unique roles in control of toxic Microcystis population and removal of MC,

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which contributes to successful transfer of toxic primary producers in aquatic food

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chains and reduction of the water risk.

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Based on aforementioned studies, control of toxic Microcystis by protozoans is a

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promising biological method, but there is only limited understanding about the

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grazing and degradation processes. Strong resistance of protozoans to MC has been

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confirmed (see Table S1), however, inner response of protozoans needs further

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investigation. Moreover, Ou et al. (2005)30 reported the efficient degradation of MC

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by flagellates possibly relied on a series of biological processes, however, the definite

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biodegradation pathways in flagellates have not been identified. These issues are

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important in enriching theoretical support in application of protozoans to control toxic

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Microcystis.

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Gene expression consists of two levels: transcription and translation.

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Transcriptomes are generally applied to analyze gene regulation network from DNA

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to mRNA, thus comparative transcriptome analysis can provide the different

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responses in gene expression level. Based on our previous significant findings that

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flagellates showed strong ability to control Microcystis and efficiently degrade

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microcystin,31,25 the present study conducted transcriptome sequencing of flagellate

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Ochromonas fed on microcystin-producing Microcystis (TM) and microcystin-free

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Microcystis (NM). The aims of this study are (1) to evaluate the transcriptomic

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responses of Ochromonas to microcystin-producing Microcystis, and (2) to identify

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potential genes and pathways involved in the resisting and degrading microcystin in

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flagellate Ochromonas. Thus, transcriptomic analyses can provide a realistic and

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intuitive explanation for the pathways of resisting and degrading microcystin as well

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as the theoretical support for control of toxic microalgae by protozoans, which may

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have a positive impact on enhancing removal efficiency of MC by protozoans through

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combining other environmental factors in applied researches.

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2. Materials and methods

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2.1 Microorganisms and culture conditions

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A clonal culture of mixotrophic flagellate Ochromonas gloeopara was isolated

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from Lake Taihu.25 To compare the effect of naturally occurring Microcystis strains on

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transcriptomic responses of Ochromonas, Microcystis aeruginosa strains FACHB-469

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and PCC7806 were used in the experiments. Both stains were originally isolated from

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natural lakes and purchased from the Freshwater Algae Culture Collection of the

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Institute of Hydrobiology, China. Microcystis strain PCC7806 can produce

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microcystin, whereas FACHB-469 is a microcystin-free strain lacking mcy gene (Fig.

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S1). The verification method of mcy D gene was provided in Supporting

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Information. All the microorganisms were maintained in sterile BG-11 medium at

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25 °C under a fluorescent light of 40 µmol photons m-2 s-1 with a 12:12 h light:dark

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cycle.

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2.2 Experimental design

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Mixotrophic O. gloeopara was fed with microcystin-producing (TM) as toxic

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treatment and microcystin-free (NM) M. aeruginosa as control in a 250-mL flask

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filled with 150 mL sterilized BG-11 medium at the conditions mentioned above.

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Correspondingly, monoculture of M. aeruginosa served as control of grazing

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treatment. The initial concentration of O. gloeopara was ~1.0×104 cells mL-1. To

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ensure O. gloeopara obtaining the same prey biomass, microcystin-producing and

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microcystin-free M. aeruginosa were fed to O. gloeopara at the same carbon content

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of 2.2 mg C L-1. Each treatment was set up in six replicates: three for growth

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experiment and the other three for RNA-Seq and RT-qPCR. The samples used in

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RNA-Seq were all taken on day 3 when the resistance and degradation processes were

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in full activity. The samples for measuring MC concentration were taken every 2 days.

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Total microcystin including intracellular and extracellular microcystin, were extracted

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according to the process used by Wilken et al. (2010),32 and then were tested

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following by ELISA kit instruction (Microcystins Plate Kit; Beacon, USA). The

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intracellular microcystin was from both Ochromonas and Microcystis. Moreover,

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photosynthetic performances (Fv/Fm, ETRmax, and αETR) of Ochromonas in NM and

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TM treatments were detected using a Phyto-PAM (Walz, Germany) at earlier stage

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(day 2, full activity in degradation process), later stage (day 6, weaker activity in

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degradation process), and final stage (day 10, MC undetectable).

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2.3 RNA isolation and sequencing

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To collect O. gloeopara cells, samples in the NM and TM groups were separately centrifuged for 15 min at 2400 g at 4 °C. Approximately 1.0×107 O. gloeopara cells

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were immediately homogenized in TransZol Up, and total RNA from the cells was

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extracted using TransZol Up Plus RNA Kit following the manufacturer’s instructions

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(ER501, TRANS, China). Then, the total RNA was purified using DNase I

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(Invitrogen, USA) to eliminate genomic DNA. The purified RNA was used in

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RNA-Seq and RT-qPCR. The concentration and integrity of RNA were verified by

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using an Agilent 2100 Bioanalyzer (Agilent, USA), and the quality was checked by

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agarose gel electrophoresis. In the study, the RNA integrity number of all samples was

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above 7.0.

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Before RNA-seq, mRNA was enriched from purified total RNA using magnetic

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beads attached with Oligo (dT), and further fragmented to synthesis cDNA as

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described by Song et al. (2017).33 Paired cDNA was repaired at the end, attached with

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Poly(A) at DNA 3’ ends, and connected sequencing adapters before being performed

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to develop a cDNA library. Finally, the cDNA library was sequenced by 1GENE

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Company (Hangzhou, China) using the Illumina HiSeq 4000 platform (Illumina,

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USA). As Microcystis is a kind of prokaryotic organism, which DNA lack Poly(A) at

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DNA 3’ ends, therefore, only O. gloeopara RNA was used in RNA-Seq in this

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experiment.

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2.4 Transcript assembly and annotation

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To obtain high-quality data, sequenced reads were cleaned up by removing reads

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with attached contaminated adapters (>5 bp bases in each read), low-quality reads

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(>20% bases of Q-score, 5%

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of nucleotides in each read). Clean reads were assembled into unigenes using De

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Novo Trinity platform (Supporting Information).34 The assembled unigenes were

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annotated by BLASTx searching in NCBI non-redundant (NR), Swiss-Prot, KEGG,

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COG, and GO databases, and mapped to NCBI Nt database by BLASTn

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(E-value1 and FDR

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≤0.001 were considered as significantly differentially expressed genes (DEGs) (Fig.

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S5). GO and KEGG analyses were also used to classify the function of DEGs.

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Pathways with Q-value≤0.05 were considered as significant enriched pathways. The

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detailed analysis method used in gene expression analysis was provided in

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Supporting Information.

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To validate the RNA-Seq data in O. gloeopara transcriptome, the expressions of

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12 DEGs mapping in all the samples were quantitated by RT-qPCR. Besides, the

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relative expressions of 5 key genes involving in degrading MC were determined at

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earlier stage, later stage, and final stage to present temporal dynamics of this process.

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Three genes as alternative reference genes were selected from unigene data by

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geNorm analysis, and α−tubulin as the most stably expressed gene was used as

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internal standard for relative expression quantification. ddH2O was used as the

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negative control in RT-qPCR. The cDNA was synthesized from mRNA by cDNA

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Synthesis SuperMix (AT311, TRANS, China), and RT-qPCR was conducted using

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TransStart Top Green qPCR SuperMix (AQ131, TRANS, China). All the primer

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sequences were presented in Table S3 under Supporting Information. Gene expression

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was calculated using 2-∆∆t method.35 Relative gene expression in temporal dynamics

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was calculated using NM treatment at each endpoint as control (setting the value to

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“1”). Correlation between RNA-Seq and RT-qPCR was performed by regression

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analysis.

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2.6 Statistical analysis Samples were collected daily to investigate population growth and every two

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days to measure the changes in microcystin. The specific growth rate (µ, d-1) of O.

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gloeopara was calculated in the following equation previously by Zhang et al. (2017)

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25

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at day a and day b. The degradation ratio of microcystin (θ, %) was calculated as

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follows25: ߠ = ሺ1 − ‫ܯ‬௧ ∕ ‫ܯ‬௖ ሻ × 100%, where Mt and Mc were microcystin

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concentrations in the grazing treatment and control.

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: ߤ ሺ݀ ିଵ ሻ = ൫݈݊஽ೌ − ݈݊஽್ ൯ ∕ ሺܽ − ܾሻ, where Da and Db were O. gloeopara densities

All data were expressed as mean ±SE in the study. Statistical analyses were

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performed using SigmaPlot 11.0.

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3. Results

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3.1 Growth of O. gloeopara and changes in microcystin concentration

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In grazing treatments, two strains of M. aeruginosa (microcystin-free and

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microcystin-producing) were both eliminated under O. gloeopara grazing, and the

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predator O. gloeopara population increased with the depletion of M. aeruginosa (Fig.

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1A). In TM grazing treatment, both intracellular and extracellular microcystin were

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finally removed (Fig. 1B). Without predator O. gloeopara, microcystin in TM control

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rose with increasing M. aeruginosa population (Figs. 1A and 1B). Moreover,

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microcystin was not detected in NM treatment. The result of PCR amplification also

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showed Microcystis strain FACHB-469 lacks mcy D gene cluster (Fig. S1). In TM

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grazing treatment, O. gloeopara grew at lower growth rate compared with the NM

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grazing treatments (Fig. 1C). In addition, the degradation ratio of microcystin was

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gradually increased and reached up to approximately 100% after 6 days, relative to

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the control of TM grazing treatment (Fig. 1D).

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3.2 Overview of assembled transcriptome

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The present study investigated the changes in O. gloeopara fed on

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microcystin-producing or microcystin-free M. aeruginosa in gene expression level by

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transcriptomic analyses. As genomic sequencing of model species in chrysophyta has

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not been conducted to date, therefore, we obtained 143779562 reads for the NM

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treatment and 164862464 reads for the TM treatment by using De Novo RNA-Seq as

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analyzed in other transcriptomic studies on O. gloeopara.36 The assembled

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transcriptome of O. gloeopara with a total of 69.18 million base pairs yielded 116969

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high-quality unigenes with a mean length of 591 bp and N50 length of 1138 bp (Table

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1). Over 85% of reads were mapped back to assembled transcriptome. Approximately

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88600 (76%), 47893 (41%), 61352 (52%), 58352 (50%), 55053 (47%), and 48148

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(41%) of these unigenes were respectively annotated through matching to Nr, Nt,

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Swiss-Prot, KEGG pathway, COG, and GO databases. Generally, a total of 94813

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unigenes (81.1%) were annotated in these public database. Regarding the species

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distribution in the Nr database, 5.8% of Ochromonas unigenes showed top matches

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with genes from Ectocarpus siliculosus, followed by Hordeum sativum (5.4%),

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Aegilops squarrosa (4.7%), Glycine max (4.4%), and Nannochloropsis gaditana (4%)

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(Fig. S2); the rest of organisms (70%) were divided into 959 species. Additionally,

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these unigenes enriched in the COG database were mainly classified into general

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function prediction only, transcription, ribosomal structure, and biogenesis and

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posttranslational modification (Fig. S3). Furthermore, these unigenes assigned to GO

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term were mainly associated with cellular process, metabolic process, cell, cell part,

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binding and catalytic activity (Fig. S4).

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3.3 Functional classification of differentially expressed genes (DEGs)

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Compared with O. gloeopara fed on microcystin-free M. aeruginosa (NM

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treatment), a total of 999 unigenes were differentially expressed in Ochromoans fed

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on microcystin-producing M. aeruginosa (TM treatment), among which 709 and 290

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genes were significantly upregulated and downregulated respectively (Figs. 2A and

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S5). All of these DEGs (999 DEGs) were used in GO (Table S4) and KEGG analysis

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(Table S5). In significantly enriched GO terms (Q-value≤0.05), different DEGs were

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categorized into several biological process, namely, translation (GO:0006412),

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protein-chromophore linkage (GO:0018289), and xyloglucan metabolic process

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(GO:0010411). Ribosome (GO:0005840), ribosomal subunit (GO:0044391), and

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cytosolic ribosome (GO:0022626) were major categories in cellular components by

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GO analysis. The main molecular function of these DEGs were structural constituent

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of ribosome (GO:0003735), structural molecule activity (GO:0005198), and

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xyloglucan:xyloglucosyl transferase activity (GO:0016762). In detail, the major GO

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terms of these upregulated DEGs were cellular process, metabolic process, organelle

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part, membrane-enclosed lumen, structural molecular activity, and binding terms (Fig.

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S6); the major GO terms of these downregulated DEGs were biological regulation,

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response to stimulus, extracellular region part, and catalytic activity (Fig. S6).

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In addition, by KEGG enrichment analysis, significantly enriched pathways

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(Q-value≤0.05) of DEGs included ribosome (89 genes, 13.6%, Ko03010),

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photosynthesis (24 genes, 3.58%, Ko00195), and starch and sucrose metabolism (24

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genes, 3.58%, Ko00500) (Table S5). Some DEGs were strongly associated with

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citrate cycle (1.34%, Ko00020), phagosome (2.53%, Ko04145), oxidative

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phosphorylation (2.98%, Ko00190), and glutamate metabolism (1.49%, Ko00250).

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3.4 Validation of gene expression in transcriptome

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To value the repeatability of the three samples under the same treatment, Pearson

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coefficient was calculated by comparing gene expressions among samples. The

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Pearson coefficients of the two treatments were all above 0.90, which conformed to

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the requirements of biological repetition. Moreover, based on the sequences obtained

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in the transcriptome, twelve differentially expressed genes (DEGs) were quantitated

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by RT-qPCR to verify RNA-Seq data. The result showed that the expression patterns

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represented a significant correlation between RT-qPCR and RNA-Seq (Fig. 2B,

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R2=0.6054, P15. Secondly, higher expression of genes

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involved in carbohydrate metabolism in the TM group included pyruvate

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dehydrogenase E1 component, phosphoglycerate kinase, and glyceraldehyde

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3-phosphate dehydrogenase in glycolysis, compared with the NM group. In pentose

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phosphate pathway, the genes encoding 6-phosphogluconate dehydrogenase and

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ribose-phosphate pyrophosphokinase were respectively 16-log2(fold change) and

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15.5-log2(fold change) higher in the TM group. In addition, most genes encoding proteins

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in citrate cycle were 17-log2(fold change) more than that in TM group. Thirdly, all the

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genes involving in oxidative phosphorylation were upregulated. Interestingly,

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mixotrophic O. gloeopara grazing on toxic M. aeruginosa obviously upregulated the

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expression of genes participating in photosynthesis and carbon fixation (Fig. 3; Table

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S6), including genes encoding Rubisco, subunits of photosynthetic systems, and

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proteins in photosynthetic electron transport. Moreover, transcriptomic analyses

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showed downregulation of auxin-responsive protein and upregulation of sphingolipid

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4-desaturase/C4-monooxygenase (Table S6).

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3.6. Notable genes related to resisting and degrading MC in O. gloeopara

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By transcriptional analysis, we found that O. gloeopara fed on toxic M.

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aeruginosa significantly upregulated the expression of genes linked to resistance (Fig.

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4; Table S6). For protein repair, the expressions of heat shock protein70 (HSP70) and

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heat shock protein 90 (HSP90) are increased. For DNA repair, gene encoding

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serine/threonine-protein phosphatase 2A catalytic subunit (PP2A) was upregulated.

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The gene expression of PP2A by RT-qPCR was higher in full activity period and

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decreased in later period in TM treatment, relative to that in NM treatment (Fig. 2C).

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Also, peroxiredoxin had a higher expression in TM group. Moreover, some signatures

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of outer protection were upregulation of O-GlcNAc transferase (OGT) and

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downregulation of chitinase by microcystin exposure.

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Pathway enrichment analysis showed phagosome was significantly enriched in

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the differentially expressed genes (Table S5). Gene encoding Rac was obviously

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upregulated (Fig. 5; Table S6). The gene expression by PT-qPCR showed Rac was

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higher in TM treatment than that in NM treatment in earlier stage (full activity in

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degrading process), and then declined in later stage when microcystin was gradually

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degraded (Fig. 2C). In addition, we found higher expression of cathepsin that mainly

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acts on hydrolysis of proteins. Glutathione S-transferase (GST), glycine

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hydroxymethyltransferase and cystathionine beta-synthase were also upregulated,

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while glutathione peroxidase was down-regulated. (Table S6).

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4. Discussion

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Ochromonas is a species of mixotrophic protists equipped with chromatoplast

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and thus classified into the Chrysophyte.37,38 To date, there is no genomic sequencing

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of model species in chrysophyta, thus O. gloeopara transcriptomic in the present

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study was De Novo assembled, which was reasonable and also used in previous

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studies of Ochromonas transcriptomics under various nutritional strategies.39,40,36

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In the long-term evolution, the ancestor of modern Chrysophyte probably has

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close relationships between those of brown algae and dinoflagellate. These organisms

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are derived from endosymbiosis of eukaryotic host cells and green algae,41 which

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could explain that assembled sequences of O. gloeopara was most hit to the model

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organism of brown algae Ectocarpus siliculosus in Nr database (Fig. S2). However,

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more than 90% of unigenes of O. gloeopara have no correlation with Ectocarpus

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siliculosus, which also indicated the diverse evolutionary trajectories in eukaryotic

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phytoplankton for the adaption to environments. Moreover, relative to a total of 94813

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unigenes annotated in Nr, Nt, Swiss-Prot, KEGG pathway, COG, and GO databases,

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the rest of 18.9% assembled unigenes was not matched to the databases mentioned

336

above (a total of 116969 high-quality unigenes assembled in the experiment). The

337

study of Lie et al. (2017)36 also showed more than 25% of genes were not annotated

338

to public databases in O. gloeopara in response to light and prey availability. Thus,

339

the function of a number of unique genes still is unknown due to lacking

340

understanding of Ochromonas genome.

341

4.1 Translation, energy metabolism, and carbohydrate metabolism

342

Enriched pathways of DEGs were translation, energy metabolism, and

343

carbohydrate metabolism (Table S6; Fig. 3). Firstly, most of the genes associated with

344

subunits in ribosome were upregulated, which had also been observed in Daphnia

345

exposed to toxic Microcystis.42,43 Protein synthesis is a significant biological process

346

supported by ribosomes. Upregulated genes in ribosome indicated that toxic M.

347

aeruginosa had significant effects on protein synthesis of O. gloeopara. Secondly,

348

higher expression of genes involved in carbohydrate metabolism in the TM treatment,

349

compared with the NM treatment. In organisms, TCA cycle is final oxidized and

350

decomposed pathways of three major nutrients (carbohydrates, lipids, and amino

351

acids), followed by only two types of production nicotinamide adenine dinucleotide

352

phosphate (NADPH) and flavine adenine dinucleotide (FADH2). NADPH belongs to

353

cytochrome P450 monooxygenase detoxification system, and it plays an important

354

role in bioconversion of toxins and maintenance of the amount of glutathione;44,45

355

FADH2 further enters into oxidative phosphorylation. In the study, some key genes in

18


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356

TCA cycle, including citrate synthase, aconitate hydratase, isocitrate dehydrogenase,

357

and malate dehydrogenase, were significantly upregulated (Table S6). In addition, the

358

gene encoding 6-phosphogluconate dehydrogenase had a higher expression level in

359

the TM treatment, which is the crucial enzyme in phosphate pentose pathway with the

360

products of NADPH and ribose.

361

As expected, all genes involved in oxidative phosphorylation were upregulated,

362

indicating that O. gloeopara enhanced the production of energy to support the above

363

metabolisms. The relative gene expression of ATP synthase (AtpB) was sharply

364

increased in full activity degrading process and decreased with decline of MC (Fig.

365

2C). Asselman et al. (2012)42 explained that the upregulation of oxidative

366

phosphorylation was intended to satisfy the additional energy requirement in Daphnia

367

exposed to stress. Interestingly, mixotrophic O. gloeopara grazing on toxic M.

368

aeruginosa obviously increased the expression of genes involved in photosynthesis

369

and carbon fixation (Table S6). Wilken et al. (2014)46 reported that the uptake of preys

370

would reduce the pigments and content of Rubisco in mixotrophic organisms. Indeed,

371

mixotrophic protists are still equipped with fully functional photosynthetic machinery

372

that serves as a reserve system for providing resources and energy after digesting

373

preys.47 Thus, O. gloeopara in the TM treatment possibly overrepresented genes in

374

photosynthesis to obtain additional organic matter by fixating carbon. Also, the

375

photosynthetic performances (Fv/Fm, ETRmax, and αETR) were higher in TM

376

treatment than those in NM treatment in full activity period (Fig. S7), in accordance

19



377

with gene expression result, which indicated energy production from photosynthesis

378

was increased during active degradation process in Ochromonas. When microcystin

379

was gradually degraded and Microcystis populations were decreased, the

380

photosynthetic parameters declined and recovered finally, which was in accordance

381

with our previous study that photosynthetic parameters decreased slightly but

382

remained at a relatively stable level with depletion of organic carbon and the

383

population reaching a stationary stage.37 Generally, enhanced ribosome and energy

384

metabolisms provided additional matter and energy for biosynthesis of proteins and

385

other matter in O. gloeopara grazing on toxic M. aeruginosa (Fig. 3). We speculated

386

that these results were closely correlated with resisting and degrading activities in O.

387

gloeopara.

388

Nevertheless, improved energy metabolism did not contribute to the growth of O.

389

gloeopara. In the present study, mixotrophic O. gloeopara grazing reduced the

390

populations of the two strains of M. aeruginosa (microcystin-free and

391

microcystin-producing) (Fig. 1A), which was in accordance with our previous

392

findings that flagellate Ochromonas can reduce Microcystis populations.25 Moreover,

393

O. gloeopara fed on microcystin-producing M. aeruginosa grew at a lower rate,

394

compared with that grown in NM treatment (Fig. 1C). Daphnia commonly decreased

395

the growth rate by toxic Microcystis exposure. Lyu et al. (2016)18 and Asselman et al.

396

(2016)43 explained that Daphnia invested energy and resources to repair misfolded

397

proteins in response to toxic Microcystis. Given these results, we speculated that O.

20


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398

gloeopara might also allocate more energy and matters in resisting and degrading

399

microcystin so that growth was reduced. Another explanation for the inhibition might

400

be other harmful substances produced by toxic Microcystis.22 Ou et al. (2005)30 also

401

reported that microcystin could damage cell structures of Poterioochromonas, thereby

402

possibly causing the death of partial cells. Moreover, a significant downregulation of

403

auxin-responsive protein and upregulation of sphingolipid

404

4-desaturase/C4-monooxygenase were detected in the study (Table S6); they are

405

commonly involved in the process of cell division and proliferation,48,49 which

406

directly confirmed the decreased growth rate of O. gloeopara in response to toxic M.

407

aeruginosa. Furthermore, the growth rate of microcystin-free strain was lower than

408

that of microcystin-producing strain in the experiment, which was different from the

409

results of previous studies,50, 51 probably due to the strain-specific difference and the

410

distinctive culturing conditions.

411

4.2 Genes involved in resisting microcystin in O. gloeopara

412

Microcystin represents a type of cyclic heptapeptides synthesized by

413

non-ribosomal pathway in Microcystis.52 Nowadays, the role of microcystin has been

414

confirmed in maintaining cyanobacterial blooms and enhancing their

415

competitiveness,53,54 while microcystin as a chemical defense are still in doubt.

416

Studies on the model aquatic animal Daphnia indicated that toxic Microcystis

417

inhibited the energy metabolism and digestion process and changed feeding

418

behavior.18,55,56 ROS induced by microcystin caused lipid peroxidation, DNA damage,

21



419

and protein damage at the cellular level of Daphnia. Lyu et al. (2014)57 reported that

420

Daphnia increased the expression of MnSOD with increased microcystin

421

concentrations.

422

Contrary to these conditions, small-sized protozoans show better tolerance to

423

toxic cyanobacteria. For instance, amoeba Naegleria can excrete toxic cyanobacteria

424

from the food vacuole, relying on food selection;58 heterotrophic flagellates

425

Diphylleia rotans can degrade cyanotoxins.27 In the present study, mixotrophic O.

426

gloeopara reduced total microcystin in the culture (Fig. 1B). Thus, we confirmed O.

427

gloeopara was equipped with degradation capacity that helps to resist toxin.

428

Moreover, results of DEGs showed a series of significantly upregulated genes in TM

429

treatment, linked with resistance (Fig. 4; Table S6). Firstly, the expressions of heat

430

shock protein70 (HSP70) and heat shock protein 90 (HSP90) are increased. HSP

431

family plays a crucial role in regulating protein modification and eliminating

432

misfolded proteins.59 Secondly, increased serine/threonine-protein phosphatase 2A

433

catalytic subunit (PP2A), a type of serine-threonine phosphorylase participating in

434

signal transport and cell apoptosis,60 might participate in repairing DNA and protein

435

damage. Microcystin usually causes histone phosphorylation by conjugating with

436

catalytic subunit of PP2A,5 thereby leading to cell apoptosis. Thus, increased

437

transcription of PP2A in the study offset the negative effects of microcystin-PP2A on

438

cells as well as promoted DNA repair. The relative gene expression result also

439

suggested PP2A was enhanced in earlier stage (full activity in degrading process) and

22


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Page 23 of 48


440

then decreased with the decline of MC (Fig. 2C), which also indicated enhanced DNA

441

repair when Ochromonas grew in full degrading activity. Thirdly, upregulated

442

peroxiredoxin could directly reduce ROS as the first line in defeating oxidative

443

stress.61 The relative gene expression of peroxiredoxin was increased in earlier stage

444

and then decreased gradually (Fig. 2C). In conclusion, O. gloeopara enhanced a series

445

of repair capacities and removal of ROS to reduce damage of microcystin to cells,

446

which was generally consistent with previous reports that Daphnia exposed to

447

microcystin improved antioxidant capacity and compensation of damage

448

proteins.18,42,43

449

Outside cells, O-GlcNAc transferase (OGT) was identified as upregulated by

450

microcystin exposure. OGT acting on the protein glycosylation often transfers

451

GlcNAc β1 to serine-threonine. Then, the proteoglycan is released and further forms

452

an extracellular matrix outside cells.62 Also, chitinase, a hydrolase of chitin, was

453

detected to be downregulated by microcystin exposure. Flagellates do not have cell

454

wall but chitin covering the cell membranes.63 Chitin is a major component of

455

epidermis in crustaceans and considered as an antioxidant involved in reducing ROS

456

and prolonging life.64 Therefore, we speculated that proteoglycan and chitin possibly

457

worked as extracellular barriers to protect O. gloeopara from dissolved microcystin

458

outside cells. Given the aforementioned consideration, O. gloeopara might strengthen

459

inner and outer strategies of protection and repair when grazing on toxic M.

460

aeruginosa and exposed to dissolved microcystin (Fig. 4).

23



461 462

4.3 Mechanisms of degrading microcystin by O. gloeopara Although some aquatic organisms resist toxic cyanobacteria to a certain extent,

463

only a few of them can biodegrade and utilize cyanotoxins. Thus, cyanotoxins are

464

generally accumulated in predators step by step, thereby inducing the increased

465

adverse impacts on the environment.20 Numerous materials have been reported to

466

reduce microcystin in waters. For instance, ferrate, hydrogen peroxide, and

467

microorganisms could degrade toxins directly;14-16 cyclodextrins could combine with

468

cyanotoxins and change their chemical structures, thereby reducing the toxicity of

469

microcystin;65 ordered mesoporous absorbed cyanotoxins to reduce the amount of

470

toxins.66 Considering that the methods of removing MC should satisfy

471

environment-friendly standards, we believe that biological degradation without

472

additional environmental burden deserves further attention. To date, various bacteria

473

have been isolated from natural waters involving cyanobacteria and identified to

474

participate in cyanobacterial lysis and removal of cyanotoxin.16 Four key enzymes

475

encoded by the mlr cluster (mlrABCD) in bacteria Sphingomonas are involved in

476

hydrolysis of circular microcystin and subsequent degradation of linear microcystin.67

477

In natural waters, the diversity of MC-degrading genotypes in the bacterial

478

community commonly shifts with the dynamics of toxic cyanobacterial blooms,68

479

because cyanotoxins are mostly stored in cells and only heavily released at the late

480

stage of blooms. In Daphnia, trypsin and ubiquitin-conjugating enzymes in intestines

481

were demonstrated to play roles in digesting microcystin.43,53

24


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Page 25 of 48


482 483

4.3.1 Hydrolytic activities in lysosomes Protozoans are not equipped with a specific digestive system. In their digestive

484

process, the preys first wrapped phagosomes and then formed food vacuoles.69

485

Subsequently, the phagosome combines with lysosome and transforms to primary

486

lysosome (Fig. 5). Thus, we speculated that the degradation process might benefit

487

from the activity in lysosome. Firstly, hydrogen peroxide (H2O2) and superoxide (O2-)

488

are major strong oxidants of clearing antigens in lysosomes as the first line of

489

resistance to pathogens, which implies that extraneous M. aeruginosa as pathogens

490

for O. gloeopara would be killed under great oxidation. Then, there are massive

491

hydrolytic enzymes involving in digesting preys in primary and secondary lysosomes.

492

Through transcriptional analysis, we observed an increased expression of cathepsin in

493

phagosome (Fig. 5 and Table S6), which is an important category of proteases

494

participating in the hydrolysis of proteins in lysosome,70 therefore, the reduction of

495

microcystin could be due to hydrolytic activities in lysosomes of O. gloeopara.

496

4.3.2 Reduction of microcystin by ROS

497

Normally, the decrease in toxicity of microcystin depends on the changes in

498

chemical structure. H2O2 and hydroxyl radical have been demonstrated to efficiently

499

degrade cyanotoxins15,71,72 because hydroxyl radical mainly attacks the sites including

500

the aromatic ring, conjugated diene in Adda, and the C=C bond in Mdha to reduce

501

microcystin.73 In lysosome, increased Rac can strongly regulate the activation of

502

NADPH oxidase to promote the production of massive ROS, including H2O2, OH-,

25



503

and O2-,74 and then H2O2 may easily be converted by catalysts to hydroxyl radical due

504

to instability of the chemical structure. In addition, increased ROS induced by toxic

505

compounds are generally harmful to organisms, thus the removal activity of ROS will

506

be enhanced. For instance, catalase (CAT) usually participates in the decomposition of

507

H2O2; glutathione peroxidase (GSH-Px) can catalyze GSH and H2O2 to generate

508

glutathione disulfide (GSSH). However, the present result showed these two genes

509

were both downregulated (Table S6). Meanwhile, the relative gene expression of Rac

510

was enhanced in full activity degrading stage and decreased when MC was gradually

511

degraded by Ochromonas (i.e. later stage and final stage) (Fig. 2C). Therefore, we

512

speculated that Rac plays an important role in Ochromoans degrading MC, and H2O2

513

was very essential for O. gloeopara fed on toxic M. aeruginosa, thereby not being

514

removed (Fig. 5). In the study, we measured microcystin concentration using ELISA

515

kit based on quantitating Adda group. The decreasing microcystin concentration

516

implied the reduction of Adda group that possibly resulted from strong oxidants

517

bonding with Adda groups. Thus, this process of O. gloeopara degrading microcystin

518

is different from that in bacteria.

519

4.3.3 Detoxication of microcystin by GSH and GST

520

Currently, glutathione S-transferase and glutathione have been believed to

521

participate in the detoxification of toxins.75 In the study, we found that some genes

522

associated with glutathione (GSH) were upregulated in O. gloeopara fed on toxic M.

523

aeruginosa, including glutathione S-transferase (GST), glycine

26


Page 26 of 48

Page 27 of 48


524

hydroxymethyltransferase and cystathionine beta-synthase (Table S6). L-cysteine and

525

glycine, which are major components of glutathione, are synthetized separately under

526

catalysis of glycine hydroxymethyltransferase and cystathionine beta-synthase (Fig.

527

5). GST functions in the cellular degradation system of toxins by catalyzing the bond

528

between glutathione (GSH) and toxins.75 Meanwhile, the relative gene expression of

529

GST by RT-qPCR was increased in full activity degrading process in Ochromonas and

530

decreased with MC being degraded (Fig. 2C). Thus, this finding was in line with

531

previous studies which reported that aquatic animals Daphnia magna and

532

Oreochromis niloticus increased the expression of GST by microcystin exposure.18,76

533

Surprisingly, temporal dynamics in expression of GST, together with the similar

534

trends in PP2A and AtpB, upregulated in the final stage, which may be a provisional

535

response to trophic conversion from heterotrophy to autotrophy of O. gloeopara after

536

M. aeruginosa was eaten up. In addition to GSH, Li et al. (2014)77 suggested cysteine

537

can also conjugate microcystin in bighead carp. Nevertheless, most of aquatic animals

538

are still unable to well resist microcystin by glutathione detoxification. It is still in

539

doubt that microcystin is as chemical defenses against metazoans. The origin of

540

microcystin is older than that of metazoans,78 which implies microcystin is not

541

directly against them. Protozoans can degrade cyanotoxins possibly because they have

542

evolved degradation mechanism during long-term grazing toxic cyanobacteria.35

543 544

Furthermore, extracellular microcystin was decreased with addition of O. gloeopara in the study. Previous study suggested filtrate of flagellates did not degrade

27



545

microcystin, which indicated the degradation process depended on intracellular

546

biological process.30 Microcystin as a polypeptide were considered to be actively

547

transported into cells,79 and the transport relied on membrane carriers on the cell

548

surface. Thus, we speculated the degradation of extracellular microcystin in O.

549

gloeopara required a specific membrane transport mechanism. At this point, further

550

clarification is needed regarding how extracellular microcystin is transferred and

551

degraded in flagellate cells.

552

In conclusion, using De Novo RNA-Seq, the present study described the

553

transcriptional regulation of mixotrophic O. gloeopara in response to

554

microcystin-producing M. aeruginosa. A total of 999 differentially expressed genes

555

were identified in comparison with TM treatment and NM treatment. These genes

556

were mostly associated with translation, carbohydrate metabolism, and energy

557

metabolism. In addition to oxidative phosphorylation, carbon fixation was enhanced

558

to provide extra resources. The excellent tolerance of O. gloeopara to microcystin

559

may be due to the strong enhancement of inner antioxidant activities and outer

560

protection. Moreover, degradation of microcystin in O. gloeopara may depend on the

561

digestion activity and the induced ROS in lysosome and GST detoxication. In this

562

context, the current study provided a better understanding of transcriptomic responses

563

of flagellates to toxic M. aeruginosa as well as highlighted the mechanisms of

564

resisting and degrading microcystin in O. gloeopara, which strongly enriched the

565

theoretical knowledge to support control of toxic microalgae by protozoans.

28


Page 28 of 48

Page 29 of 48


566

Supporting information

567

Supporting Information Method describes details on models and software used in

568

gene expression analysis, PCR amplification, and agarose gel electrophoresis method

569

(PDF).

570

Supporting Information Figure (PDF).

571

Supporting Information Table describes Table S1 (A list of protozoans that can ingest

572

Microcystis and resist or degrade MC) and Table S2 (the settings used in Trinity to

573

generate the De Novo assembly) (PDF).

574

Table S3 describes primer sequences employed in RT-qPCR (XLSX).

575

Table S4 describes enriched GO terms for mixotrophic Ochromonas fed on

576

toxin-producing and microcystin-free Microcystis (XLSX).

577

Table S5 describes enriched KEGG pathways for mixotrophic Ochromonas fed on

578

toxin-producing and microcystin-free Microcystis (XLSX).

579

Table S6 describes differentially expressed genes of mixotrophic Ochromonas

580

correlating to energy metabolism, translation, carbohydrate metabolism, resisting and

581

degrading microcystin in the form FPKM (Fragments Per kb per Million fragments).

582

(NM: microcystin-free Microcystis; TM: microcystin-producing Microcystis)

583

(XLSX).

584

Acknowledgments

585

This study was supported by National Natural Science Foundation of China

586

(31730105, 31870444), Major Project of Natural Science Research for Universities in

29



587

Jiangsu Province (17KJA180007), the Priority Academic Program Development of

588

Jiangsu Higher Education Institutions, and the Topic Selection of Excellent Doctoral

589

Dissertations of Nanjing Normal University (1812000006385).

590

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591

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detoxication. Biochim. Biophys. Acta 1998, 1425, 527-533.

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41



839

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840

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841

79. Eriksson, J. E.; Grönberg, L.; Nygård, S.; Slotte, J. P.; Meriluoto, J. A. O.

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843

Biochim. Biophys. Acta 1990, 1025, 60-66.

844

42


Page 42 of 48

Page 43 of 48


845

Table 1. Summary of transcriptome dataset of mixotrophic Ochromonas. (NM:

846

microcystin-free Microcystis; TM: microcystin-producing Microcystis)

NO. of reads

NM

TM

143779562

164862464

NO. of nonredundant contigs

116969

Transcriptome Size

69.18 Mb

Contig N50

1138 bp

Mean length

591 bp

Reads mapped back to assembly

85.60%

847

43


87.20%


848

Fig. 1. (A) Population dynamics in NM and TM treatments. (B) Changes in

849

microcystin concentration in TM groups. (C) The specific growth rates of

850

mixotrophic Ochromonas in NM and TM groups. (NM treatment: microcystin-free

851

Microcystis with or without Ochromonas; TM treatment: microcystin-producing

852

Microcystis with or without Ochromonas). (D) The degradation ratio of microcystin

853

(%).

854

44


Page 44 of 48

Page 45 of 48


855

Fig. 2. (A) Numbers of differentially expressed genes (DEGs) between TM and NM

856

treatments. 290 down-regulated genes and 709 up-regulated genes were in TM

857

treatment, compared with NM treatment. (B) Correlation between RNA-Seq and

858

RT-qPCR data. 12 differentially expressed genes were identified using RT-qPCR. (C)

859

The heatmap of relative gene expression of Rac, GST, PP2A, PER, and AtpB.

860

“Control” represented NM treatment. Sample1, Sample2, and Sample3 represented 3

861

replicates in treatments. T1, T2, and T3 respectively represented earlier stage (day 2,

862

full activity in degrading process), later stage (day 6, weaker activity in degrading

863

process), and final stage (day 10, MC undetectable). (Rac: Ras-related C3 botulinum

864

toxin substrate; GST: glutathione S-transferase; PER: peroxiredoxin Q; PP2A:

865

serine/threonine-protein phosphatase 2A catalytic subunit; AtpB: ATP synthase CF1

866

beta-subunit) A

C

800

709

Number of DEGs

down-regulated up-regulated

T1

600

Rac 400 290

GST

200

0

RNA-Seq (log2 of fold change)

B

PER TM vs NM

20

PP2A

19 18

AtpB 17 16

0

5

10

T3

Relative expression (TM/NM)

2

R = 0.6054 P < 0.001

2-11 20

15

867 868

T2

Control Sample1 Sample2 Sample3 Sample1 Sample2 Sample3 Sample1 Sample2 Sample3 Sample1 Sample2 Sample3 Sample1 Sample2 Sample3

15

RT-qPCR (log2 of fold change)

45


22

213


Page 46 of 48

869

Fig. 3. Schematic model illustrating the global changes of cellular metabolic pathways.

870

Details about these genes are provided in Supplementary Table S6. (1.

871

Phosphoglucomutase; 2. Fructokinase; 3. GAPDH; 4. Phosphoglycerate kinase; 5.

872

Pyruvate dehydrogenase E1 component; 6. Citrate synthase; 7. Aconitate hydratase; 8.

873

Aconitate hydratase; 9. Isocitrate dehydrogenase; 10. Isocitrate dehydrogenase; 11.

874

Malate dehydrogenase; 12. UTP-glucose-1-phosphate uridylyltransferase; 13.

875

6-phosphogluconate dehydrogenase; 14. Ribose-phosphate pyrophosphokinase; 15.

876

NADH dehydrogenase; 16. Cytochrome c reductase; 17. Cytochrome c oxidase; 18.

877

ATP synthase; 19. Ribulose-bisphosphate carboxylase; 20. Photosystem II P680

878

reaction center D1 protein; 21. Cytochrome b6; 22. Plastocyanin; 23. Photosystem I

879

P700 chlorophyll a apoprotein A1; 24. Ferredoxin; 25. Ferredoxin--NADP+ reductase;

880

26. F-type H+-transporting ATPase)

NADPH

1

α-D-Glucose-1P

12

α-D-Glucose-6P

Chloroplast

D-Gluconate β-D-Fructose-6P

2

6-Phospho-D-gluconate

3

H2O

D-Ribulose-5P

4 Glycerate-3P

PSII Cytochrome b6/f complex

14

Acetyl-CoA

5-Phosphoribosyl diphosphate

Pyruvate

26

ATP

Fd 24 22 PC

O2

NADP+

25 FNR

21

D-Ribose-5P

5

19

NADPH

20

13

Glycerate-1,3P2

β-D-Fructose

Calvin ADP CO2 cycle 3P-glycerate

UDP-glucose

PSI 23

6

O2

cis-Aconitate Complex I

9

Citrate

6

Complex IV

8 Isocitrate

7

TCA cycle

Oxalosuccinate

H+

15

11

2-Oxo-gultarate

ATP Complex V

NAD+

10

Oxaloacetate

NADH/FADH2

e+ Q

16

17 e+

e+

Cytochrome bc1 Complex III

(S)-Malate

Mitochondrion NADPH

Ribosome

18 H 2O

FADH2

881 882

46


Proteins Large subunit Small subunit Amino acids

Page 47 of 48


883

Fig. 4. The mechanism for resisting microcystin in mixotrophic Ochromonas

884

transcriptome. (ROS: reactive oxygen species; HSP: heat shock protein; PP2A:

885

serine/threonine-protein phosphatase 2A catalytic subunit)

886 887

47



888

Fig. 5. Putative mechanism for mixotrophic Ochromonas degrading microcystin. The

889

dashed box indicated the process that Microcystis was digested and MC was degraded

890

in Ochromonas. (MC: microcystin; GSH: glutathione; GST: Glutathione S-transferase;

891

GSH-Px: glutathione peroxidase; GSSH: glutathione disulfide; CAT: catalase)

892 893 894

48


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