An in vitro and in vivo study of broad-range phosphopantetheinyl

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An in vitro and in vivo study of broad-range phosphopantetheinyl transferases for heterologous expression of cyanobacterial natural products Tianzhe Liu, Rabia Mazmouz, and Brett A. Neilan ACS Synth. Biol., Just Accepted Manuscript • DOI: 10.1021/acssynbio.8b00091 • Publication Date (Web): 21 Mar 2018 Downloaded from http://pubs.acs.org on March 22, 2018

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ACS Synthetic Biology

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An in vitro and in vivo study of broad-range phosphopantetheinyl transferases for heterologous expression of cyanobacterial natural products Tianzhe Liu†, Rabia Mazmouz†‡, and Brett A. Neilan*† ‡ †

School of Biotechnology and Biomolecular Sciences, The University of New South Wales, NSW 2052, Sydney, Australia ‡ School of Environmental and Life Sciences, The University of Newcastle, NSW 2308, Callaghan, Australia * Corresponding author. E-mail: [email protected]

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Abstract

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Phosphopantetheinyl transferases catalyze the post-translational modification of

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carrier proteins involved in both primary and secondary metabolism. The functional

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expression of polyketide synthases and non-ribosomal peptide synthetases requires

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the activation of all carrier protein domains by phosphopantetheinyl transferases. Here

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we describe the characterization of five bacterial phosphopantetheinyl transferases by

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their substrate specificity and catalytic efficiency of four bacterial carrier proteins.

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Comparative in vitro phosphopantetheinylation analysis showed Sfp possesses the

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highest catalytic efficiency over various carrier proteins. In vivo co-expression of

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phosphopantetheinyl transferases with carrier proteins revealed a broad range

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substrate specificity of phosphopantetheinyl transferases; all studied

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phosphopantetheinyl transferases were capable of converting apo- carrier proteins,

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sourced from diverse biosynthetic enzymes, to their active holo form.

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Phosphopantetheinyl transferase co-expression with the hybrid non-ribosomal peptide

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synthetases / polyketide synthases mcy gene cluster expression confirmed that the

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higher in vitro activity of Sfp translated in vivo to a higher yield of production.

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Table of Contents Graphic:

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Keywords: phosphopantetheinyl transferases, heterologous expression, natural

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products, microcystin, non-ribosomal peptide synthetase, polyketide synthase, NRPS,

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PKS, biosynthetic gene cluster

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Phosphopantetheinyl transferases (PPTases) are a superfamily of enzymes required

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for the post-translational modification of carrier proteins (CPs) from fatty acid

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synthases (FASs) involved in primary metabolism, or polyketide synthases (PKSs)

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and non-ribosomal peptide synthetases (NRPSs) involved in secondary metabolism 1.

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PPTases catalyze the transfer of a phosphopantetheinyl (Ppant) moiety from

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coenzyme A (CoA) to a conserved serine residue in CPs, converting the enzyme from

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an inactive apo- form to an activated holo- form 2. Based on catalytic specificity, PPTases can be divided into two families in

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prokaryotes 3: The first group is AcpS-type PPTase, which is associated with primary

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metabolism and typically modify acyl carrier proteins (ACP) from FASs 2. The

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second group of PPTase, typified by Sfp, has broader substrate specificity, and

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modify CPs not only from FASs but also from PKSs and NRPSs 1, 4. Cyanobacteria have been recognized as prolific producers of natural products

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possessing unique structures and diverse bioactivities 5-7. The fact that cyanobacteria

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are either slow growing and/or uncultivable is a hindrance to the large-scale

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production of cyanobacterial natural products 5, 8. The lack of tools for genetic

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manipulation of this group of organisms also hampers the elucidation of natural

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product biosynthesis pathways. To overcome these obstacles, heterologous expression

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in a genetically amenable and fast-growing host has become a new means to

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characterize pathways and achieve optimal production of their secondary metabolites

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9, 10

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renders this organism a desirable expression host 11, 12, with many bioactive natural

. The short doubling time and ample genetic toolbox available for Escherichia coli



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products having been heterologously produced in this organism 9, 13-19. The coding

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gene for a PPTase is not always clustered with a biosynthesis gene cluster, and E. coli

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PPTases show limited activity in modifying CPs from secondary metabolites 20-22. For

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these reasons it is necessary to introduce a PPTase into this heterologous host to

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facilitate activation of a broad range of cyanobacterial CPs.

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Previous studies of cyanobacterial PPTases mainly focused on their involvement

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in the activation of their cognate pathways represented by a glycolipid biosynthesis in

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heterocyst-forming cyanobacteria 23, 24. Insufficient studies have been conducted

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exploring the potential of cyanobacterial PPTases to activate non-cognate CPs.

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NsPPT, formerly referred to as PPTNs, the Sfp-type PPTase from Nodularia

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spumigena NSOR10 was the first cyanobacterial PPTase shown to possess broad

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substrate specificity by efficiently modifying non-cognate CPs 25, and for a long time

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(2007-September 2017) being the only cyanobacterial PPTase which is able to

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activate broad natural product biosynthetic pathways. This PPTase was proven to not

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only efficiently modify the PKS-ACP in glycolipid synthase from Nostoc punctiforme

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(ArCPNp), but also act as a strong catalyst for activating non-cognate PKS-ACP

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(NosB-ACP, formerly referred to as ACPNp 25) and NRPS-PCP (McyG-PCP); being

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associated with nostopeptolide and microcystin synthase/synthetase in Nostoc

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punctiforme ATCC 29133 and Microcystis aeruginosa PCC 7806, respectively.

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Despite being structurally similar to Sfp, the PPTase from Synechocystis sp. PCC

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6803, Sppt, does not possess broad-range specificity over non-cognate CPs involved

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in secondary metabolism 26. Two other Sfp-type cyanobacterial PPTases, SePPT


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(from Synechococcus elongatus PCC 7942) and FPPT (from Fischerella sp. PCC

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9339), failed to modify both MACP (a PKS ACP from M. aeruginosa NIES 843) and

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APNPCP (a NRPS PCP from Anabaena sp. PCC 7120) 27. The biased catalytic

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activity of Sppt 26, SePPT 27 and FPPT 27 toward non-cognate cyanobacterial CPs

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indicates that cyanobacterial PPTases are not always the preferred catalysts for

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activating cyanobacterial natural products biosynthetic pathways. Hence, it can be

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difficult to assess the substrate range of a PPTase from structural analysis alone. Thus

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it is worthwhile to study the catalytic characters of PPTases from the other organisms

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toward the modification of cyanobacterial CPs.

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Here we present the study of the relative affinity of five diverse bacterial PPTases

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to modify four bacterial CPs from various biosynthetic pathways. To characterize the

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ability of these PPTases to activate NRPS/PKS CPs, both in vivo and in vitro studies

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were conducted, elucidating both their catalytic specificity and efficiency on the CPs

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and their application in heterologous expression of cyanobacterial natural products.

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The in vitro assay revealed the efficiency of the PPTases by measuring the conversion

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rate of CPs from unmodified apo- form to modified holo- form. Furthermore, via co-

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expression of the PPTase and CP, we were able to assess the capability of the tested

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PPTase to activate cyanobacterial CPs from various biosynthetic pathways. Finally, to

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assess their suitability for heterologous expression of cyanobacterial biosynthetic

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pathways, the hybrid NRPS-PKS synthetase, responsible for microcystin biosynthesis,

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was assessed by comparing the yields of microcystin obtained from co-expression

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with the various PPTases.



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RESULTS AND DISCUSSION

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In silico analysis of PPTases and CPs

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A multiple alignment of five PPTase sequences was performed using T-Coffee and

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ESPript (Figure 1). The PPTases show low identity (16.9 to 29.2%) but a higher

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similarity (35.9% to 64.6%), with the exception of the two cyanobacterial PPTases

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(NsPPT and MaPPT) which share 42.2% identity (Supporting Information Tables S1

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and S2). Sfp (type II PPTase), being the most structurally and biochemically

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characterized PPTase 4, 23, 28, 29, was used as a reference for comparisons in this study.

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The multiple alignment showed that the main amino acids involved in the

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active site of Sfp are conserved within all five sequences (Figure 1). Most notable was

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the conservation of the acidic residue E151 (Sfp was used as the reference for amino

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acids numbering), which is hypothesized to facilitate transfer of the Ppant arm via

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deprotonation of the hydroxyl group of the catalytic serine in CP followed by a

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nucleophilic attack of the β-phosphate group of CoA 28. Residues binding to the CoA

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moieties, including G74, K75 and P76 were well conserved within all PPTases,

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except Svp that had a substitution (K75A) 23. Interestingly, residues binding CoA’s

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adenosine-phosphate (T44, K28 and K31) were not conserved. The CoA α-phosphate

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is linked to Sfp via residues S89, H90 and K155 that are well conserved in other

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PPTases, with the exception of Svp which shows a polymorphism at this position

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(S89T). Residues involved in the binding of the essential Mg2+ cofactor are D107,

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E109 and E151 which are conserved in all sequences 29.


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Two interaction sites between CP and Sfp have been identified. The first site

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(Y36) is conserved in MtaA; whereas the second site of contact is a hydrophobic

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patch conserved among all the sequences. To compare the tertiary structures of

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selected PPTases, the SWISS-MODEL server was used to infer the three-dimensional

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structures of Svp, MtaA, NsPPT and MaPPT. The X-ray structure of Sfp and

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predicted three-dimensional structures of Svp, MtaA, NsPPT and MaPPT are

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compared in Figure 2. The predicted structures of MtaA, NsPPT and MaPPT have the

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same secondary structure and topology as Sfp. The C-terminal helix, however, could

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not be predicted (Figure 2). Svp shows a predicted structure with less secondary

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structures, including the first helix which could not be modelled along with the three

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β-strands 7, 8, 11 and the helix 7.

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Figure 1. Sequence alignment of PPTases from diverse organisms. Sfp from B. subtilis, Svp from S. verticillus, MaPPT from M. aeruginosa PCC 7806, NsPPT from N. spumigena NSOR10,



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and MtaA from S. aurantiaca. Sequence alignment created using T-coffee and ESPript. White characters with red background indicates 100% identity, red characters with a blue frame indicate similarity.

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Figure 2: Three-dimensional structure prediction of five PPTases using SWISSMODELING. (A) Sfp. (B) Svp. (C) MtaA. (D) NsPPT. (E) MaPPT.

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Four cyanobacterial CPs were selected for alignment and prediction of their

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three-dimensional structures (Figures 3 and 4). CPs were chosen for this study due to

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their involvement in the biosynthesis of cyanobacterial toxins, the most intensely

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studied cyanobacterial natural products, which impact the environment and human

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health. CP sequences show very low identity (12.2% to 25.4%), but with higher

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similarity (30.6% to 48.7%) (Supporting Information Tables S3 and S4). The

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alignment shows the catalytic serine residue bearing the Ppant arm is conserved in all

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CPs (Figure 3). Furthermore, the DSL triad is conserved in NosB-ACP, CyrB-ACP,

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SxtA-ACP, but is replaced by HSL (commonly found in CPs sequences) in McyG-

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PCP 30-32. The glycine residue (-3 from the catalytic serine residue), necessary for

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orienting the CP for PPTase interaction, is conserved in all CPs 31, 32. The two amino

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acids involved in the direct interaction with the PPTase are a conserved Leu (+1 from

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catalytic serine) and a hydrophobic residue (Val, Leu or Ile) located +4 from the

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active site. The X-ray structure of McyG-PCP and homology models of NosB-ACP,


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CyrB-ACP, SxtA-ACP are compared in Figure 4. The four homology models show

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the topology of all CPs, three main α-helices and a small helix (Figure 4) 31, 32.

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The structure of Sfp in complex with CoA and TycC3-PCP was used as a model

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to predict the interactions between the different combinations of PPTases and CPs

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(Supporting Information Figure S1). In all complexes, the CoA has been proposed to

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be trapped between the PPTases and CPs. The interaction between CoA and PPTases

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is conserved in the presence or absence of CPs and mainly involves K28, K31, Y73,

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G74, K75, P76, S89, H90, D107, E109, E151 and K155 in the Sfp-CoA-TycC3-PCP

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complex 28, 31, 32. This site is well conserved amongst the different combinations of

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PPTase-CP interactions (Supporting Information Figures S1 and S2). The distance

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between S57 in CyrB-ACP, CoA and the catalytic residue E151 of PPTase is longer

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than the other complexes (Supporting Information Figure S3). The interactions

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between PPTases and CPs, the hydrophobic pocket and the hydrogen bonds, are also

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conserved (Supporting Information Figures S4 and S5). The best fit to the model was

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seen with the complexes NosB-ACP and SxtA-ACP, whereby K73, L79 for NosB-

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ACP and D58, L64 for SxtA-ACP, reside within the hydrophobic pocket of the

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PPTase. In contrast, the complexes with CyrB-ACP show weaker interactions

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(Supporting Information Figures S4 and S5). For example, T52 does not lie in the

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hydrophobic pocket and the hydrogen bond between CyrB-ACP and PPTases is 3.5

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Å, too long for it to be considered optimal 33. Conversely, the optimal H-bond (2.6-2.7

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Å) was observed for the complexes with McyG-PCP. In general, the H-bonds have



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been described as mostly non-covalent weak bonds with a donor-acceptor distances

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between 2.5-3.2 and 3.2-4 Å 33.

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Figure 3. Sequence alignment of carrier proteins. McyG-PCP from microcystin synthetase, CyrB-ACP from cylindrospermopsin synthase, SxtA-ACP from saxitoxin synthase, and NosBACP from nostopeptolide synthase. Sequence alignment was made by using T-coffee and ESPript. White characters with red background shows 100% identity, red characters with blue frame indicate similarity.

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Figure 4: Three-dimensional structure prediction of four CPs using SWISS-MODELING. (A) CyrB-ACP. (B) SxtA-ACP. (C) NosB-ACP. (D) McyG-PCP.

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Phosphopantetheinylation of non-cognate carrier proteins in vitro

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To compare catalytic efficiency, both PPTases and PKS ACP- and NRPS PCP-

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domains were expressed in E. coli after induction, followed by the affinity

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purification (Supporting Information Figures S6 and S7).

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Time coursed phosphopantetheinylation assays monitored the rate of CP conversion from apo- to holo- form catalyzed by PPTases, which was quantified by


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HPLC. As shown in the Supporting Information (Figures S8-S11), apo- CPs in the

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presence of PPTase and CoA were converted to the holo- form and eluted earlier than

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the unmodified CPs from the affinity column. This allowed us to quantify the amount

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of CP converted to the holo- form in a specific incubation time, and elucidate the

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catalytic efficiency of PPTase over specific substrate. Remarkable differences in the

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catalytic efficiency of PPTases over CPs were seen, with the exception of SxtA-ACP.

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As shown in Figure 5, Sfp exhibited the best in vitro phosphopantetheinylation

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activity, with 83% CyrB-ACP, 55% NosB-ACP and 75% McyG-ACP being modified

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in the five-minute assay. Although MtaA efficiently modified McyG-PCP and NosB-

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ACP (with 86% and 26% of the CPs, respectively, modified in five minutes), its

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activity on CyrB-ACP was not as efficient as that of the other PPTases in this study.

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Notable catalytic bias was also detected for the other three PPTases during the

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modification of certain CPs. When NsPPT was assayed for McyG-PCP modification,

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only 18% CP was converted to the active holo- form in the 30-minute reaction.

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Similarly, low efficiency was evident in the reaction between Svp and NosB-ACP

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(18% conversion in 30 minutes) and, surprisingly, modification of McyG-PCP by

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MaPPT (3% conversion in 30 minutes).

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Figure 5. Relative catalytic efficiency of PPTase in vitro. (A) CyrB-ACP. (B) SxtA-ACP. (C) NosB-ACP. (D) McyG-PCP.

224 Catalytic efficiency is a critical criterion for a PPTase to be useful for

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heterologous production of natural products. A previous study revealed that Svp has

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remarkably higher activity with its cognate ACP (TcmM) compared with the

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exogenous PPTase Sfp 34. This further prompted studies on PPTases for the

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production of related secondary metabolites, such as erythromycin 35. Cyanobacterial

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PPTases have previously been shown to prefer their cognate CPs as substrates, while

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exhibiting limited activity in modifying non-cognate CPs 27. Our current study,

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however, revealed that the cognate PPTase does not always exhibit superior CP

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modifying activity compared to PPTases from other organisms. This is demonstrated

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by MaPPT being the least efficient catalyst for conversion of its cognate CP McyG-

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PCP. Similarly, neither NsPPT or MaPPT were the optimal catalyst for any of the four

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cyanobacterial CPs. As mentioned previously, the cyanobacterial Sfp type PPTase-

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Sppt lacks the capacity to modify the cyanobacterial CPs McyG-PCP and NosB-ACP

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, which also suggests that PPTase substrate specificity/catalytic efficiency is not


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necessarily in accordance with its species phylogeny. Considering that horizontal gene

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transfer influences the evolution of biosynthetic gene clusters 36, the resulting

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interspecies shuffling of coding genes may render co-occuring CPs and PPTases as

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less efficient phosphopantetheinylation pairs 37.

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The in vitro catalytic efficiency of PPTases also varies between different CPs.

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Although Sfp shows a high efficiency with all CPs studied, the other PPTases studied

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exhibited a degree of bias for certain CPs. Of the PPTases used in this study, MtaA

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was the best catalyst for McyG-PCP but was the least efficient for CyrB-ACP; NsPPT

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exhibited strong efficiency for activation of NosB-ACP, as for MaPPT with CyrB-

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ACP, while these two cyanobacterial PPTases had minimal catalysis of McyG-PCP.

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This remarkable shift in catalytic efficiency between proteins from taxonomically

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related bacterial sources emphasizes the importance of using a broad-range PPTase

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for carrier protein modification.

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The comparison between the predicted data from the three-dimensional structure

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modeling and the in vitro phosphopantetheinylation results shows that in silico

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prediction is helpful if caution is applied to the data. Globally the prediction shows

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that the structure of different PPTases are highly similar; the residues involved in

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catalysis or interaction (with CoA and/or CP) are well conserved, with the main

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exception being two substitutions in the Svp sequence on positions involved in

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binding with CoA. This could provide an explanation of the low efficiency of Svp in

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the in vitro assay. The prediction of CP structure showed that there are no major

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structural differences between all CPs used in this study. The predicted data for



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PPTase-CoA-CP complexes show that all the combinations are similar, however

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small differences do exist; for example, SxtA-ACP and NosB-ACP show the best fit

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of all the CPs for the hydrophobic pocket of all PPTases. This prediction reflects the

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successful conversion to holo-SxtA-ACP for all PPTases. NosB-ACP had a moderate

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conversion rate, but is far from that of SxtA-ACP.

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The strength of the hydrogen bond between McyG-PCP and the PPTases was

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predicted to be the strongest of all the CPs, however, this does not appear to have a

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considerable effect on the rate of conversion as McyG-PCP shows one of the lowest

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conversion rates. The predicted structure for CyrB-ACP in complex presented a weak

270

hydrogen bond with PPTases, with the CP exhibiting a poor fit for the hydrophobic

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pocket of the PPTases. Contrary to the prediction, CyrB-ACP showed moderate

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conversion rates in the in vitro assay, comparable to NosB-ACP.

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Phosphopantetheinylation of non-cognate carrier proteins in vivo

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To test the effectiveness of the PPTases for the in vivo activation of different CPs,

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their coding genes were cloned and co-expressed in E. coli. MALDI-TOF/TOF mass

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spectrometry was used to detect a 340 Da mass shift, corresponding to the addition of

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the Ppant moiety from CoA to the conserved catalytic serine residue in the CPs. As

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shown in Figure 6 and Supporting Information (Figure S12), this mass shift was

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detected among all the CPs co-expressed with PPTases, confirming the in vivo

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phosphopantetheinylation activity of all PPTases by NosB-ACP, CyrB-ACP, SxtA-

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ACP, and McyG-PCP.


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284 285 286 287

Figure 6. MALDI-TOF/TOF of in vivo phosphopantetheinylation of CPs. (A) CyrB-ACP. (B) SxtA-ACP. (C) NosB-ACP. (D) McyG-PCP. (1) CP expressed alone. (2) CP co-expressed with MaPPT.

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The PPTases had varied in vivo phosphopantetheinylation capacity. Incomplete

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modification of NosB-ACP was observed from co-expression with Sfp and Svp (apo-

291

NosB-ACP ~20% and 5%, respectively, Supporting Information Figure S13) while

292

NosB-ACP co-expressed with other PPTases resulted in full conversion to holo-

293

NosB-ACP.

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Interestingly, we also observed that the PPTase of E. coli was able to

295

phosphopantetheinylate SxtA-ACP, with approximately 30% of this CP produced in

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E. coli present as the active holo- form in the absence of a Sfp-type PPTase. As the

297

majority of this CP remained in its apo- form, a Sfp-type PPTase needs to be co-

298

expressed to yield a fully functional CP, reinforcing the need for an exogenous PPTase

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for heterologous expression in the E. coli system.

300 301

As shown by phosphopantetheinylation experiments, non-cognate CPs from diverse biosynthetic pathways are able to be modified, both in vitro and in vivo, by



302

PPTases of cyanobacterial origin (NsPPT and MaPPT) and from other diverse phyla,

303

namely the Gram-positive bacterium Bacillus (Sfp), Actinobacteria (Svp) and

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Myxobacteria (MtaA). Our results demonstrate the suitability of the studied PPTases

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for producting a broad range of cyanobacterial natural products in E. coli.

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In vivo activation of a multi-domain NRPS-PKS gene cluster

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A competent PPTase for heterologous expression should avoid bias for a certain type

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of CP and efficiently catalyze all CPs of a given pathway, since NRPS and PKS

310

biosynthetic functionality requires all CPs to be fully modified by the PPTase. Thus,

311

assessing the in vivo post-translational modification of a gene cluster comprising both

312

NRPS and PKS is critical for the evaluation of the catalytic efficiency of the PPTases.

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We previously reported the heterologous expression of the 55 kbp microcystin

314

biosynthetic gene cluster (mcy) encoding for a hybrid NRPS-PKS, including seven

315

PCPs and four ACPs 18. By co-expressing the mcy gene cluster with various PPTases,

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the yield of the cyclic pepetide was explored using a scale of induction controlled by

317

IPTG concentration. Microcystin yield changed based on the IPTG concentration

318

(Figure 7). High levels of induction (50 µM and 100 µM of IPTG) of all PPTases,

319

with the exception of Sfp, resulted in a significant decrease in [D-Asp3]microcystin-

320

LR production, represented by a 60% and 95% yield losses, respectively, compared

321

with the low induction group (10 µM IPTG). MtaA, NsPPT, and MaPPT showed

322

similar trends with similar microcystin yields under low induction levels, representing

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the optimal yields using these PPTases. Although Svp showed similar production


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patterns, a notable increase of yield (84%) under 10 µM IPTG induction compared

325

with the non-induced counterpart was indispensable. Sfp expression was the least

326

sensitive to induction levels, showing high tolerance to intense IPTG induction with

327

the optimal microcystin yield occurring under 50 µM IPTG, and similar yields under

328

0 µM and 100 µM IPTG. PPTase concentration under 0-100 µM of IPTG induction

329

was investigated (Figure 8). This confirmed that the levels of protein did increase with

330

higher levels of inducer, however there was no significant difference in PPTase

331

concentrations between 10 and 50 µM IPTG. Most notably, leaky expression from the

332

T7 promoter 38 produced a high basal level of PPTase production, equating to more

333

than half of the concentration produced under 100 µM IPTG.

334

335 336 337 338 339 340

Figure 7. Yield of [D-Asp3]microcystin-LR in E. coli BL21(DE3). pFos-PbiTet-mcy was coexpressed with different pET-PPTase plasmids with varied IPTG induction intensity for in vivo CP phosphopantetheinylation (*P < 0.05 and ** P < 0.01, analysis was performed using Student’s t test in GraphPad Prism 6). [D-Asp3] microcystin-LR was not detected in the control sample (recombinant BL21(DE3) carrying the empty pET vector).

341



342 343 344 345 346

Figure 8. PPTase concentration in E. coli BL21(DE3). PPTase co-expressed for CP phosphopantetheinylation with varied IPTG induction concentration applied for PPTase production. (*P < 0.05 and ** P < 0.01, analysis was performed using Student’s t test in GraphPad Prism 6).

347 348

Unlike expressing a single CP where the modification efficiency might not be an

349

issue, the functional expression of large multi-module synthetases/synthases requires

350

extra considerations for PPTase selection. As demonstrated here, higher yields of

351

microcystin were produced when Sfp was used for phosphopantetheinylation of CPs

352

in the microcystin synthetases/synthases. This result suggests a higher in vivo catalytic

353

activity of Sfp in modifying carrier proteins, which is consistent with the in vitro

354

phosphopantetheinylation assay.

355

Previous studies have shown that the yield of the desired secondary metabolites

356

could be manipulated by over-expressing or knocking out relevant PPTase(s). As

357

prolific producers of specialized metabolites, the majority of these studies were

358

performed in Actinomyces. For example, overproduction of SchPPT in Streptomyces

359

chattanoogenisis L10 increases natamycin yield by about 40% 39. A single/double

360

PPTase(s) knockout in Streptomyces coelicolor, a strain natively containing three

361

promiscuous PPTases, produces both yield increase and decrease of certain secondary

362

metabolites 40. Zhang et al. also recently showed that yield increase caused by PPTase


Page 18 of 32

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363

over-expression contributes not only to the production optimization, but also the

364

discovery of new metabolites by increasing their production to a detectable level 37.

365

Our current study demonstrates that the production of secondary metabolites in E.

366

coli is also modulated by PPTase activity, as microcystin yields were altered under

367

different PPTase expression levels (Figure 7). Low levels of PPTase (10 µM IPTG

368

induction) provided the highest levels of microcystin yields, which was particularly

369

evident for Sfp and Svp. However, Sfp was the only PPTase to also produce high

370

yields with 50 µM IPTG induction. Given that all CPs studied were fully converted to

371

the holo form in vivo, the observed differences are unlikely to be an effect of

372

incomplete activation of the synthetase, but may be more reflective of increased stress

373

upon the host from high PPTase production.

374

We propose that this could be explained as a ‘saturation of CP modification’

375

model, resulting in poor megasynthetase functionality. That is, supplementation of

376

PPTase improves the production titer of secondary metabolites when the PPTase is

377

undersupplied, by facilitating complete modification of apo- carrier proteins. Once the

378

PPTase concentration has reached saturation, any further increase of PPTase levels

379

may adversely affect biosynthesis of the pathway via occlusion of the CP domain.

380

This “saturation model” may apply to the incompletely modified NosB-ACP from

381

nosB-ACP-sfp co-expression construct (Figure S13). To completely modify all carrier

382

proteins an increase of the relative expression level of Sfp over NosB-ACP may be

383

required, or via extension of the post-induction time as described previously 41.

384



Page 20 of 32

385

CONCLUSIONS

386

In summary, we showed that the optimal concentration of PPTase varies with each

387

enzyme, and is likely related to the efficiency of CP interactions during

388

phosphopantetheinylation. Moreover, the broader the PPTase substrate specificity, the

389

less sensitive NRPS-PKS functionality is to the concentration of PPTase. This

390

emphasizes the importance of not only selecting a suitable PPTase for heterologous ,

391

but also the suitable induction conditions that complement NRPS/PKS expression

392

levels. In characterizing these PPTases, our study enriches the genetic tools that can

393

be used in synthetic biology, facilitating the discovery of natural products encoded by

394

novel or cryptic gene clusters.

395 396

METHODS

397

Table 1. PPTases and CPs used in this study. Protein

PPTase Sfp Svp

MtaA NsPPT MaPPT

Source organism

Comments

NCBI Accession

Protein ID

Ref.

CP011051.1

AJW85878.1

4

AF210311.1

AAG43513.1

34

CP002271.1

ADO71799.1

42

AY646183.1

AAW67221.1 24

AM778958.1

CAO88702.1

43

microcystin gene cluter AM778952.1 (mcy) cylindrospermopsin gene EU140798.1 cluster (cyr)

CAO90231.1

43

ABX60161.1

44

Bacillis subtilis subsp. Gram-positive bacteria, spizizenii str. W23 firmicutes Streptomyces actinobacteria verticillus ATCC 15003 Stigmatella aurantiaca myxobacteria DW4/3-1 Nodularia spumigena cyanobacteria NSOR10 Microcystis cyanobacteria aeruginosa PCC 7806

CP McyG-PCP M. aeruginosa PCC7806 CyrB-ACP Cylindrospermopsis raciborskii AWT 205


Page 21 of 32 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


SxtA-ACP NosB-ACP

Cylindrospermopsis raciborskii T3 Nostoc punctiforme ATCC 29133

saxitoxin gene cluster (sxt) EU629178.1

ACF94636.1

45

nostopeptolide gene cluster (nos)

WP_0124087 84.1

25

CP001037.1

398 399

In silico analysis of proteins used in this study

400

The PPTases and CPs used in this study were sourced from diverse microorganisms

401

(Table 1). The sequences used in this study were aligned by T-Coffee and analyzed by

402

ESPript. The identity and similarity matrices were generated using the online software

403

SIAS (http://imed.med.ucm.es/Tools/sias.html). The prediction of secondary

404

structures was done with PSIPRED (http://bioinf.cs.ucl.ac.uk/psipred/) and adjusted

405

manually with the data from three-dimensional structure modelling. Three-dimensional structure modelling was performed using SWISS-MODEL (46-

406 407

48

408

solved structure of Sfp crystalized with CoA and TycC3-PCP (S45A) (PDB ID:

409

4mrt). The template used to model McyG-PCP was the McyG-A-PCP (PDB ID:

410

4r0m), for CyrB-ACP the CP domain from MLSA2 of the mycolactone polyketide

411

synthase (PDB ID: 5hv8), for SxtA-ACP the PfACP monomer (PDB ID: 3gzm), and

412

for NosB-ACP the CP domain from holo-AB3403 a four domain non-ribosomal

413

peptide synthetase (PDB ID: 4zvh). Prior to modelling, the SWISS-MODEL server

414

was tested for the de novo prediction of Sfp structure and McyG-PCP (see Supporting

415

Information Figure S14). All structures were visualized by Yasara View software

416

(http://www.yasara.org/products.htm#view) to generate all figures presented in this

417

study, and to predict both the hydrogen bonds and the distance inter-atoms.

, https://swissmodel.expasy.org). The template for PPTase modelling was the X-ray



418 419

Construction of expression plasmid

420

All PPTase and CP coding genes were amplified using Velocity polymerase (Bioline)

421

following the manufacturer’s recommendations, and purified with DNA Clean and

422

Concentrator-5 kit (ZymoResearch). The PCR products for sfp maPPT, nsPPT, mtaA

423

were digested with NcoI and XhoI (New England Biolabs), svp was digested with

424

NdeI and XhoI, then purified and ligated into pET28b (Novagen) that was previously

425

digested using the same restriction enzymes and gel purified (Gel DNA Recovery Kit,

426

ZymoResearch). CP genes were cloned into pET28b vector using a similar strategy

427

(NcoI and XhoI digestion for cyrB-ACP and sxtA-ACP, NdeI and XhoI for mcyG-

428

PCP). All genes were amplified from their corresponding genomic DNA, with the

429

exception of nosB-ACP which was cloned previously 24.

430

The plasmid used for in vivo phosphopantetheinylation was constructed by

431

deleting His-tag associated with the PPTase gene, based on the method described by

432

Hemsley et al 49, followed by cloning of the His-tagged CP coding genes into the

433

pET28b-PPTase plasmids. The construction was completed by classical cloning with

434

the insertion of His-tagged CP amplicons at BglII site. An extra helix (coloured in

435

grey as shown in Figure 4 (D)) was added to McyG to obtain more stable expression.

436

All primers used in this study are listed in Supporting Information (Table S5).

437

Protein expression and purification

438

The constructed plasmids were expressed in E. coli BL21 (DE3) containing the

439

plasmid pRARE 50. With 1% overnight culture inoculation, cells were cultured in


Page 22 of 32

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440

terrific broth (TB) with 50 µg.mL-1 kanamycin and 34 µg.mL-1 chloramphenicol, at

441

37°C under 200 rpm agitation until an absorbance of 0.6-0.8 at 600 nm. To express all

442

recombinant proteins used in this study (PPTases, CPs, and PPTase-CPs co-

443

expression), the cells were induced with 200 µM isopropyl β-D-thiogalactoside

444

(IPTG) and incubated at 18°C overnight before harvesting by centrifugation for 20

445

min at 3,220 × g (Eppendorf 5804R) and frozen at -20°C.

446

The cell pellets were thawed on ice, and resuspended in lysis buffer (20 mM

447

NaH2PO4, 500 mM NaCl, 20 mM imidazole, 10% glycerol, pH 8.0) and lysed on ice

448

by sonication at 30% amplitude for 2 min (1 s pulse followed by 4 s pulse off,

449

Branson 450 digital sonifier with a 3 mm probe). Cellular debris was pelleted by

450

centrifugation at 48,000 × g for 30 min at 4°C (Hitachi CR-GIII). The desired protein

451

was purified from the soluble fraction by using a HiTrap chelating column (GE

452

Healthcare) with a linear gradient of imidazole from 20 mM to 500 mM in 50 mL (20

453

mL from 20 mM to 120 mM, 10 mL isocratic when reached 120 mM imidazole, and

454

20 mL from 120 mM to 500 mM). All fractions were analyzed on denaturating

455

polyacrylamide gel electrophoresis (SDS-PAGE). The purest protein-containing

456

fractions were pooled and desalted using Amicon centrifugal units (10 kDa size for

457

PPTases and 3 kDa size for CPs, Millipore), then frozen in liquid nitrogen and storage

458

at -80°C. Protein concentration was measured by a Bradford protein assay kit (Bio-

459

Rad). SDS-PAGE of all proteins used in this study after desalting are shown in the

460

Supporting Information (Figures S6 and S7).

461



462

In vitro phosphopantetheinylation assay and detection of

463

phosphopantetheinylation by high performance liquid chromatography (HPLC)

464

Pantetheinylation assays were adapted from previously studies 1, 24. In brief, the

465

reaction mixtures (100 µL) containing 50 mM HEPES (pH 7.4), 150 mM NaCl, 10

466

mM MgCl2, 500 µM CoA, 5 mM Tris(2-carboxyethyl)phosphine (TCEP), 50 µM

467

carrier protein, and 500 nM PPTase were incubated at 37°C for 5, 10, 30 min, and

468

terminated by adding 10 µL of 500 mM EDTA.

469

The relative catalytic efficiency of PPTases was analyzed with the HPLC

470

method by measuring the conversion of apo-CP to holo-CP 51. Reaction mixture (100

471

µL) was injected into an analytical HPLC column (Jupiter 5u C18 300A column,

472

Phenomenex Australia Pty Ltd) equilibrated with 40% acetonitrile in 0.1%

473

trifluoroacetic acid (TFA). A linear gradient with buffer A: 0.1% formic acid in water,

474

buffer B: acetonitrile at 1 mL min-1 was used to elute the carrier protein with constant

475

column temperature of 27.5°C (method for CyrB-ACP and SxtA-ACP: Time 0

476

min=5% B, 5 min=32% B, 10 min=37% B, 50 min=47% B, 55 min=52% B, 57

477

min=95% B, 59 min=95% B, 61 min=5% B, 71 min=5% B; method for NosB-ACP

478

and McyG-PCP: Time 0 min=5% B, 5 min=37% B, 10 min=42% B, 20 min=44.5%

479

B, 22 min=47% B, 24 min=95% B, 26 min=95% B, 28 min=5% B, 33 min=5% B);

480

the absorbance at 220 nm was monitored to analyze the elution profile. Holo-CP

481

eluted faster than apo-CP under these conditions, and the conversion rate of apo- to

482

holo- can be measured by comparing the peak area of these two forms of CP with the

483

software Chemstation (Agilent).


Page 24 of 32

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484 485

Detection of in vivo phosphopantetheinylation by MALDI-TOF/TOF mass

486

spectrometry

487

To evaluate the specificity of the PPTases, the masses of target carrier proteins were

488

detected by MALDI-TOF/TOF mass spectrometry. For matrix preparation, 10 mg of

489

3,5-dimethoxy-4-hydroxyl cinnamic acid was added into 1 mL 80% acetonitrile with

490

0.1% TFA, and 1 mL of matrix was mixed with 1 µL of protein sample (1 mg.mL-1)

491

on the surface of a MALDI target plate, followed by analysis by Bruker

492

ultrafleXtreme MALDI-TOF/TOF with a YAG laser. Data acquisition was performed

493

in the positive ion mode and the instrument calibrated immediately prior to each

494

analysis. Analysis was performed in the linear delayed extraction mode acquiring 100

495

averaged spectra.

496 497

In vivo activation of multi-domains NRPS-PKS gene cluster

498

A constructed plasmid containing microcystin (mcy) biosynthetic gene cluster (pFos-

499

biTet-mcy) 18 was used to conduct the PPTase in vivo CP-activation analysis of a

500

multi-domain NRPS-PKS. Before co-expression, the kanamycin (KanR) and

501

chloramphenicol (CmR) resistance cassettes were replaced with a spectinomycin

502

resistance marker (SpecR) on plasmid pFos-biTet-mcy as shown in Supporting

503

Information (Figure S15). E. coli strain BL21 (DE3) was transformed with the

504

generated fosmid via electroporation, and the clones were selected on LB agar plates

505

containing 20 µg.mL-1 apramycin and 50 µg.mL-1 spectinomycin, to generate BL21-



506

pFos-biTet-mcy-specR. The pET28b-PPTase plasmids were subsequently chemically

507

transformed into BL21-pFos-biTet-mcy-specR to generate BL21-pFos-biTet-mcy-

508

specR-PPTase (Sfp/Svp/MtaA/NsPPT/MaPPT) selected by 20 µg.mL-1 apramycin, 50

509

µg.mL-1 spectinomycin and 50 µg.mL-1 kanamycin.

510

Previously described protocols for recombinant cell fermentation, microcystin

511

extraction and quantification 18 were adopted for this study, albeit with modification

512

of the antibiotics used for selection. The production of PPTases, driven by T7

513

promoter, was induced with 0 µM, 10 µM, 50 µM or 100 µM of IPTG when 0.5

514

µg.mL-1 of tetracycline induction was applied for transcription of mcy genes.

515 516

ASSOCIATED CONTENT

517

Supporting information

518

This material is available free of charge via the Internet at http://pubs.acs.org.

519

Primer sequences, identity and similarity for the PPTases and CPs, SDS-PAGE of

520

purified PPTases and CPs, predicted 2D and 3D-structures, sequence alignment of

521

CPs, in vitro phosphopantetheinylation of CP monitored by HPLC, in vivo

522

phosphopantetheinylation of CPs monitored by MALDI-TOF/TOF, Yield of [D-

523

Asp3]microcystin-LR in E. coli BL21(DE3).

524 525

ABBREVIATIONS


Page 26 of 32

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526

PPTase: phosphopantetheinyl transferase; NRPS: non-ribosomal peptide synthetase;

527

PKS: polyketide synthase; PCP: peptidyl carrier protein; ACP: acyl carrier protein;

528

FAS: fatty acid synthase; CoA: coenzyme A; IPTG: isopropyl β-D-thiogalactoside.

529 530

AUTHOR INFORMATION

531

Corresponding Author

532

* E-mail: [email protected]

533

Authors' contributions

534

T.L., R.M. and B.A.N. designed the overall project. T.L. performed the primary

535

sequence alignment, plasmid construction, protein expression and purification, in

536

vitro and in vivo phosphopantetheinylation assay, fermentation, extraction and

537

quantification of microcystin with the technical support of R.M. R.M. performed the

538

secondary and third-dimensional structure prediction as well as the active sites

539

prediction of all proteins. T.L., R.M. and B.A.N. analysed that data and wrote the

540

manuscript.

541

Notes

542

The authors declare no competing ﬁnancial interest.

543 544

ACKNOWLEDGMENTS

545

The authors would like to thank Chris Marquis and Helene Lebhar for the access to

546

their protein purification platform at UNSW, Anne Poljak and Sydney Liu Lau for

547

assisting MALDI-TOF/TOF analysis, Sohail Siddiqui and Sarah Ongley for



548

proofreading the manuscript. This research was supported by the Australian Research

549

Council Linkage Project grant LP140100642 and Diagnostic Technology P/L. T.L.

550

was funded by the China Scholarship Council (CSC).

551


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An in vitro and in vivo study of broad-range phosphopantetheinyl

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