Transcriptome analysis guided metabolic engineering of Bacillus subtilis for riboflavin production

ARTICLE IN PRESS Metabolic Engineering 11 (2009) 243–252

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Regular Article

Transcriptome analysis guided metabolic engineering of Bacillus subtilis for riboflavin production Shuobo Shi a,b,1, Tao Chen a,b,1, Zhigang Zhang c,d, Xun Chen a,b, Xueming Zhao a,b, a

Department of Biochemical Engineering, School of Chemical Engineering and Technology, Tianjin University, Tianjin 300072, China Key Laboratory of Systems Bioengineering, Ministry of Education, Tianjin University, Tianjin 300072, China c BioTechnology Institute, 1479 Gortner Avenue, University of Minnesota, St. Paul, MN 55108, USA d Department of Chemical Engineering and Materials Science, 421 Washington Avenue SE, University of Minnesota, Minneapolis, MN 55455, USA b

a r t i c l e in fo

abstract

Article history: Received 24 December 2008 Received in revised form 20 April 2009 Accepted 5 May 2009 Available online 13 May 2009

A comparative transcriptome profiling between a riboflavin-producing Bacillus subtilis strain RH33 and the wild-type strain B. subtilis 168 was performed, complemented with metabolite pool and nucleotide sequence analysis, to rationally identify new targets for improving riboflavin production. The pur operon (purEKBCSQLFMNHD) together with other PurR-regulated genes (glyA, guaC, pbuG, xpt-pbuX, yqhZ-folD, and pbuO) was all down-regulated in RH33, which consequently limited the supply of the riboflavin precursors. As 5-phospho-ribosyl-1(a)-pyrophosphate (PRPP) strongly inhibits the binding of PurR to its targets, it was inferred that the reduced expression of PurR-regulated genes might be caused by a low PRPP pool, which was subsequently confirmed by metabolite analysis. Thus, we selected and cooverexpressed prs and ywlF genes in RH33, which are involved in the biosynthetic pathway of PRPP from ribulose-5-phosphate. This co-amplification led to an elevated PRPP pool and thus the increased transcript abundances of PurR-regulated genes participated in riboflavin precursor biosynthesis. The riboflavin titer was increased by 25% (up to 15 g l1) in fed-batch fermentation. & 2009 Elsevier Inc. All rights reserved.

Keywords: Riboflavin Bacillus subtilis Transcriptome PRPP pool Metabolic engineering

1. Introduction In the biotechnology industry, classical strain improvement has achieved a long history of success (Crueger and Crueger, 1984; Peberdy, 1985), which relied on random mutagenesis and selection. These methods are still very useful, especially with the development of efficient strain selection methods (Gall et al., 2008). However, unwanted changes in physiology and growth retardation may occur alongside the desired improvements. Since the introduction of metabolic engineering, a more rational improvement approach emerges for microbial development (Bailey, 1991; Stephanopoulos et al., 1998). Riboflavin (vitamin B2) serves as a precursor for the synthesis of the coenzymes flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD), which are used as electron acceptors for many oxidoreductases (Stahmann et al., 2000). As such, riboflavin is required for a wide variety of cellular processes and is supplemented for feed and food fortification purposes in humans and animals to maintain health. The Gram-positive bacterium Bacillus  Corresponding author at: Department of Biochemical Engineering, School of Chemical Engineering and Technology, Tianjin University, Tianjin 300072, China. Fax: +86 22 27406770. E-mail address: [email protected] (X. Zhao). 1 These authors contributed equally to this work.

1096-7176/$ - see front matter & 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.ymben.2009.05.002

subtilis, currently the most competitive riboflavin producer, has been widely adopted in the commercial riboflavin production processes (Knorr et al., 2007; Wu et al., 2007; Zamboni et al., 2003). The biosynthesis of riboflavin in B. subtilis occurs through seven enzymatic steps starting from GTP and ribulose-5-phosphate, which is shown in Fig. 1. The riboflavin producer B. subtilis was initially developed using the ‘‘classical’’ strain development approach that relied on iterative cycles of random mutagenesis and selection to create genetic diversity and identify improved riboflavin mutants (Perkins et al., 1991; Stahmann et al., 2000). Then, a number of conceivable strategies were carried out to construct high-level riboflavin-producing B. subtilis strains. These include enhancement of both gene dosages and transcriptional level of riboflavin operon in the mutants (Perkins et al., 1999), constitutive expression of the key gene (ribA) in riboflavin + biosynthetic pathway (Humbelin et al., 1999), enhancing energy generation and reducing maintenance metabolism (Zamboni et al., 2003), increasing precursor supply by modulating carbon flow through pentose phosphate pathway (Zamboni et al., 2004a; Zhu et al., 2006), and deregulating gapB expression by ccpN knockout based on screening B. subtilis transposon mutants (Ta¨nnler et al., 2008). However, these strategies were not based on a comprehensive analysis of the complex microbial metabolism and regulation, which would further facilitate the successful and efficient

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Fig. 1. Schematic overview of expression profiles of genes involved in relevant pathways of riboflavin production. The numbers are the ratios of the comparative expression levels in B. subtilis strain RH33 vs. 168. Blue lettering indicates downregulation, red indicates up-regulation, and black indicates without notable changes. The genes that were underlined were selected for overexpression. G6P, glucose-6-phosphate; F6P, fructose-6-phosphate; GAP, D-glyceraldehyde 3-phosphate; PGA, 3-phosphoglycerate; PYR, pyruvic acid; AcoA, acetyl coenzyme A; Ser, serine; Gly, glycine; 10-formyl-THF, 10-formyl tetrahydrofolate; gluconate-6-P, 6-phospho-D-gluconate; Ru-5-P, ribulose-5phosphate; Ribo-5-P, ribose-5-phosphate; PRPP, phosphoribosylpyrophosphate; Gln, glutamine; X5P, xylulose 5-phosphate; E4P, D-erythrose 4-phosphate; IMP, inosine monophosphate; XMP, xanthosine monophosphate; GMP, guanosine mono-phosphate; GTP, guanosine tri-phosphate; DARPP, 2,5-diamino-6-ribosylamino-4(3H)pyrimidinone-50 -phosphate; ARPP, 5-amino-6-(50 -phosphoribosylamino)uracil; ArPP, 5-amino-6-(50 -phosphoribitylamino)uracil; ArP, 4-(1-D-ribitylamino)-5-amino-2,6dihydroxypyrimidine; DHPB, 3,4-dihydroxy-2-butanone 4-phosphate; DRL, 6,7-dimethyl-8-ribityl-lumazine; FAD, flavin adenine dinucleotide (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.).

microbial metabolic engineering (Nielsen and Olsson, 2002). Recently, as a holistic and discovery-driven approach, transcriptome analysis has been successfully applied in metabolic engineering for rationally probing the complex gene and metabolic regulatory networks. Novel target genes have been identified to optimize microbial production strains (Choi et al., 2003; Harris et al., 2009; Izallalen et al., 2008; Jaluria et al., 2007; Park et al., 2007; Peebles et al., 2009; Sindelar and Wendisch, 2007; Wierckx et al., 2008). Here we took a strategy to increase riboflavin production in a riboflavin-producing B. subtilis strain based on comparative transcriptome analysis between a riboflavin high-producer RH33 and wild type 168, integrated with DNA sequencing and metabolite pool measurement. Our integrated approach allowed us to understand genome-wide transcriptional differences underlying strain performance, and to identify potential metabolic bottlenecks for riboflavin production. We rationally overexpressed two genes simultaneously to activate precursor purine biosynthesis pathways by modulating global regulator activity via metabolite pool manipulation. This strategy was capable of elevating riboflavin titer by 25% in B. subtilis RH33 (up to 15 g l1) in fed-batch fermentation.

2. Materials and methods

Table 1 Strains and plasmids used in this study. Strain or plasmid Strains B. subtilis 168 B. subtilis RH33 B. subtilis RH33-Prs B. subtilis RH33-PY E. coli Top10 Plasmids pUC18 pSG1192 pHPL10 pRPU10 pRPU12 pRPU13 pRPU14 pRPU15

Description of genotype

Source

Wild-type

BGSC

r

r

Em , Cm , containing multiple riboflavin operons Emr, Cmr, Spcr, containing a P43-prs (CDS) fragment integrated in the chromosome of RH33 Emr, Cmr, Spcr, containing a P43-prs-ywlF (CDS) fragment integrated in the chromosome of RH33 Host strain for constructing plasmids

Laboratory stock This study This study Laboratory stock

Ampr Ampr, Spcr pHP13, containing a P43 promoter Ampr, Ampr, Ampr, Ampr, Ampr,

BGSC BGSC Laboratory stock containing a prs (CDS) fragment This study containing a P43-prs (CDS) fragment This study Spcr, containing a P43-prs (CDS) fragment This study containing a P43-prs-ywlF (CDS) fragment This study Spcr, containing a P43-prs-ywlF (CDS) fragment This study

BGSC, Bacillus Genetic Stock Center (http://www.bgsc.org/).

2.1. Strains and plasmids Bacterial strains and plasmids used in this work are listed in Table 1. Two different riboflavin-producing mutants B. subtilis RH13 and B. subtilis J617 were derived from B. subtilis 168 by multiple rounds of selection with azaguanine (Azr), decoyinine (Dcr) and roseoflavin (RoFr) for resistance mutations that deregulate the riboflavin biosynthetic pathway. An integration vector pRB63 was constructed by inserting a native rib operon into

pDG364 (BGSC, http://www.bgsc.org/). Then the vector pRB63 was integrated into the B. subtilis RH13 chromosome at the native rib operon locus by single crossover and selected at 5 mg ml1 chloramphenicol. Subsequently, the copy number of the integrated pRB63 was increased by selecting colonies that grew at higher chloramphenicol concentrations. Colonies that were able to grow up with 40 mg ml1 chloramphenicol were isolated, and one was denoted as B. subtilis RH13H[pRB63]n. Another

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0.03 g l1 ZnCl2  7H2O, 0.04 g l1 MnCl2  4H2O. The cells were grown in the batch mode for 6 h after inoculation, and then the feeding process was continued from 6 to 48 h. The feed solution contained 650 g l1 glucose, 10 g l1 yeast extract, 10 g l1 NaNO3, 5 g l1 NH4NO3, 5 g l1 KH2PO4, 5 g l1 K2HPO4, and 2 g l1 MgSO4  7H2O. The residual glucose concentration in the fermentor was maintained no more than 10 g l1 by controlling the glucose feed rate manually. The cultivation was carried out for 48 h at the agitation rate of 900 rpm, with an aeration rate of 1.1 vvm and temperature of 41 1C. The head pressure is maintained at 0.5 atm. pH was maintained at 6.8 with 1 M H2SO4 and 10% ammonia. All the experiments were carried out independently in biological triplicates, and the reported results were the average of three replicate experiments.

integration vector pRB62 carried erythromycin resistant was integrated into the B. subtilis J617 chromosome at the native thrC locus by double crossover. The vector was constructed by insertion of a 4.4-kb P43-modified rib operon (substituting the native ribP1 promoter and RFN regulatory element with the constitutive P43 promoter) into pDG1664 (BGSC, http:// www.bgsc.org/). The resulting strain was denoted as B. subtilis J617H[pRB62]. Subsequently, B. subtilis RH13H[pRB63]n and B. subtilis J617H[pRB62] were fused by protoplast fusion. The fusants were selected in solid regeneration medium containing 40 mg ml1 chloramphenicol and 5 mg ml1 erythromycin. One clone was denoted as B. subtilis RH33, which was used as the parental strain in our study. Plasmids were constructed by the following procedures. Firstly, polymerase chain reaction (PCR) was used to amplify structural genes of prs and ywlF with the following primers: prs-up 50 GGGGCCCGGGCCAGAGCGAGACAAGTAAA, prs-down 50 -GGGGGAGCTCGCTAGCTCCTATTACAAACAATACCCA, ywlF-up 50 -GGGGCCCGGGGCTAGCGGCTGCGCGGTCAATA, ywlF-down 50 -GGGGGAGCTCGCGGCCGCTTGTTTCAATTCCGCTTGGTC, based on the published B. subtilis 168 genome sequence (Kunst et al., 1997). The prs PCR product was ligated into pUC18 using XmaI and SacI restriction sites to construct pRPU10. The constitutively expressed P43 promoter (Wang and Doi, 1984) was obtained from the vector pHPL10 (unpublished work) after digested with BamHI and XmaI. It was then ligated in the BamHI and XmaI restriction sites of pRPU10 to get plasmid pRPU12. The ywlF PCR product was digested with NheI and SacI, and cloned into NheI-SacI-sites of pRPU12 to construct pRPU14. To facilitate further research, a spectinomycin resistance gene (spc) from pSG1192 was inserted at the SalI site of pRPU12 and pRPU14 to give plasmids pRPU13 and pRPU15, respectively. All DNA manipulations were carried out as described previously (Sambrook and Russell, 2001). LB medium supplemented with corresponding antibiotics was used as the standard medium for plasmid construction in Escherichia coli. The plasmids were transformed into competent cells of B. subtilis RH33 as described previously (Anagnostopoulos and Spizizen, 1961).

About 25 ml of the culture was used for preparation of total RNA, which was extracted using RiboPureTM-Bacteria Kit (Ambion, UK). Total RNA was stored at 70 1C until use. The quantity and quality of RNA was analyzed by UV spectrophotometry and denaturing formaldehyde agarose gel electrophoresis, respectively. Probe sets on the B. subtilis Genome Array (Antisense) were designed based on the wild-type B. subtilis 168 genome sequence data of Kunst et al. (1997) by Affymetrix (Santa Clara, CA, USA). An aliquot of 10 mg B. subtilis total RNA was used to synthesize first-strand cDNA with random primers and SuperScript II reverse transcriptase. Then the cDNA was fragmented to 50–200 bp and labeled by biotin. After hybridization at 45 1C for 16 h at 60 rpm, the microarray slides were washed and stained on Affymetrix Fluidics Station 450. The scanned images were obtained with the GeneChip Scanner 3000 (Affymetrix) and were analyzed using the default setting of GeneChip Operating Software (GCOS 1.4). Then a LOWESS normalization method was performed to normalize the different arrays using dChip software. The differentially expressed genes were identified through overlapping gene analysis of biological duplicate experiments using a 2-fold change as an empirical criterion.

2.2. Growth conditions

2.4. Quantitative RT-PCR

For transcriptome and quantitative RT-PCR analyses, 5-phospho-ribosyl-1(a)-pyrophosphate (PRPP) pool, and riboflavin measurements, a preculture of B. subtilis RH33 and B. subtilis 168 were inoculated from a fresh LB agar plate and cultured to exponential growth period in LBG medium (LB medium with 1% glucose) at 37 1C. The intracellular metabolites, PRPP, ribose-5-phosphate and ribulose-5-phosphate, were determined during exponential growth period in minimal medium with 20 g l1 glucose, 2 g l1 (NH4)2SO4, 13.1 g l1 KH2PO4, 6 g l1 K2HPO4, 1.2 g l1 NaC6H5O7  2H2O, 10 mg l1 MgSO4  7H2O and supplemented with tryptophan, phenylalanine and tyrosine (25 mg l1 each). Fed-batch fermentation was performed in B. subtilis strains for riboflavin overproduction. The strain was revived by growing on LB agar slants. The seed cultures of the revived B. subtilis strains were prepared in 500-ml shake flasks at 41 1C with 240 rpm. The seed medium contained 20 g l1 glucose, 5 g l1 yeast extract, 10 g l1 NaNO3, 5 g l1 NH4NO3, 1 g l1 KH2PO4, 1.65 g l1 K2HPO4, 1.5 g l1 MgSO4  7H2O, 0.03 g l1 FeCl2, 0.04 g l1 MnCl2  4H2O, 0.04 g l1 ZnCl2  7H2O. After 14 h incubation at 41 1C, 150 ml of the seed culture was transferred into in a 5-l bioreactor (Bao Xing, China) containing 2350 ml fermentation medium. The initial medium of fed-batch fermentations contained 20 g l1 glucose, 5 g l1 yeast extract, 10 g l1 NaNO3, 5 g l1 NH4NO3, 4 g l1 KH2PO4, 7.5 g l1 K2HPO4, 1.5 g l1 MgSO4  7H2O, 0.03 g l1 FeCl2,

To validate transcriptome results, relative abundances of selective transcripts were measured by quantitative reverse transcription-PCR, which was carried out by an iCycler (Bio-Rad, USA) using the QuantiTect SYBR Green RT-PCR kit (Qiagen, Germany) according to the manufacturer’s instructions. In brief, 100 ng of DNA-free total RNA was used in a total reaction volume of 50 ml with 0.4 mM of each primer (Supplementary Table 1). The fold change of each transcript in each sample relative to the control sample was measured in triplicates, normalized to internal control gene gapA and calculated according to the comparative Ct method (Livak and Schmittgen, 2001).

2.3. Transcriptome analysis

2.5. Metabolite concentration measurement Immediately upon harvest, 5 ml of cell sample was added to 15 ml of quenching fluid containing 70 mM HEPES in 60% (v/v) aqueous methanol. The samples were centrifuged to separate the quenching fluid and the remaining cell pellets were resuspended in 35% (v/v) perchloric acid. After one freeze–thaw cycle the sample was neutralized by 5 M K2CO3. The precipitate was removed by another centrifugation, and resulting supernatants were stored at 80 1C until analysis. Measurement of intracellular PRPP concentrations was carried out using the methods described by Kornberg et al. (1955). The ribose-5-phosphate and ribulose-5-phosphate

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were measured using a Luminescence Spectrophotometer (Hoque et al., 2005).

Table 2 Validation of microarray data by quantitative RT-PCR. Genes

Quantitative RT-PCR

Microarray

purF purD glyA folD prs ywlF ribA narG

0.2570.01 0.1170.01 0.4270.02 0.4470.02 0.9570.02 0.8770.01 15970.37 2.9970.05

0.3870.01 0.5070.01 0.6870.03 0.6070.04 1.0070.03 0.9070.06 13.2770.08 6.4870.06

2.6. Enzyme activity assay in crude extracts The crude cell extract was prepared as described previously (Fisher and Mangasanik, 1984). For enzyme assay, strains were cultivated at 37 1C in minimal medium. Quantification of PRPP synthetase activity, ribose-5-phosphate isomerase activity, and transketolase activity were performed as described (Hove-Jensen and Maigaard, 1993; Iida et al., 1993; Sakuma et al., 1991). Specific activity was expressed as nmol min1 mg1 protein. Protein content was determined according to the Bradford method. 2.7. Analytical methods Cell growth (OD600 nm) was monitored with a spectrophotometer. Cell growth rate was calculated by log-linear regression analysis of OD600 nm versus time. Glucose concentration was determined using a glucose analyzer (Model-SBA40, Shandong, China). For riboflavin measurements, the samples were diluted with 0.05 M NaOH to the linear range of the spectrophotometer, and centrifuged at 12,000 rpm for 2 min to remove the cells. Then the OD444 nm was immediately measured. The culture broth was centrifuged and the resulting supernatant was used for measurement of the concentrations of extracellular metabolites (acetate, acetoin, etc.) by using a HPLC system (Agilent, HP1100, USA) equipped with a UV–visible light monitor (Bai et al., 2004). 2.8. Nucleotide sequencing Selected genes involved in riboflavin biosynthesis and cellular central metabolism were sequenced for both regulatory and coding regions, using the traditional Sanger method by Beijing AuGCT biotechnology Co., Ltd., China.

3. Results and discussion 3.1. Comparative transcriptome analysis Transcriptome profiles of a riboflavin-producing B. subtilis RH33 was compared to wild-type B. subtilis 168 on DNA microarray using mid-exponential phase cell samples (OD600 nm0.90) grown in LBG medium. In B. subtilis RH33, this condition corresponded to the state of high riboflavin production, which was coupled to cell growth. The specific growth rate of RH33 and 168 were 1.5770.15 and 1.6170.15 h1, respectively. During mid-exponential phase, the specific growth rates were nearly constant, without discernible accumulation of byproducts such as acetate (data not shown). Our past experiences showed that there was no significant difference in transcriptional profiles at different time-points of mid-exponential phase. The microarray experiment was very reproducible. The Pearson correlation coefficient between the biological duplicates was about 0.95; the overlapping of differentially expressed genes identified using an empirical 2-fold criterion from each experiment was more than 90%. It was found that 619 genes showed significant variations at the genome-wide transcriptional level between RH33 and 168, representing about 15% of B. subtilis genome. Quantitative RT-PCR experiment was performed in parallel to validate microarray data (Table 2). The average fold change of ribA and narG transcripts, were identified to be 159 and 2.99 in RH33 relative to 168, respectively. This was consistent with microarray data, since both genes were significantly up-

Fold change of each transcript in B. subtilis RH33 relative to 168 was reported.

regulated to 13.27 and 6.48-fold, respectively. The genes prs and ywlF were identified not to be differentially expressed (1.00 and 0.90) by microarray, which was confirmed by quantitative RT-PCR (0.95 and 0.87). Furthermore, by quantitative RT-PCR, the mean expression levels of purF, purD, glyA, and folD were identified to be 0.25, 0.11, 0.42, and 0.44, respectively. All the four genes were identified as significantly down-regulated (0.38, 0.50, 0.68, and 0.60) in B. subtilis RH33 by transcriptome analysis. Therefore, the correlation between quantitative RT-PCR and microarray was good (Table 2), suggesting the validity of the microarray gene expression measurements. The differentially expressed genes fell into nearly all functional categories (Supplementary Table 2). The largest group with altered transcriptional levels in the B. subtilis RH33 was a group of 83 genes encoding for the transport/binding proteins and lipoproteins, all of which are associated with the cell membrane. Forty-eight affected genes belonged to the group involved in metabolism of amino acids and related molecules, especially aspartate, cysteine, glutamate, histidine, leucine, threonine, tyrosine, and serine biosynthesis genes. Other large functional groups changed significantly included 24 genes associated with motility, 23 genes associated with carbohydrate metabolism (notably myoinositol and acetoin metabolism), 23 genes associated with metabolism of nucleotides and nucleosides (mainly purine biosynthetic genes), 32 genes associated with transcription regulation and 27 genes associated with phage-related function. Therefore, it seems that complex transcriptional regulation mechanism(s) underpinned the strain performance differences, directly or indirectly affecting riboflavin production. Some of these transcriptional effects will be discussed in detail as follows.

3.2. Nucleotide sequencing of selected genes Since B. subtilis RH33 was developed mostly by repeated random mutagenesis and selection, mutations that affect gene function were also expected to occur. The identification of mutational changes is necessary to fully understand riboflavin synthesis processes in RH33. However, the unknown random mutations may affect the activity of gene product, but may not affect gene expression per se and thus such mutations may not be detected by microarray. To identify possible beneficial mutations for riboflavin production in RH33, we selected 67 genes for DNA sequencing, including both regulatory and coding regions. These genes participated in riboflavin biosynthesis, precursor supply and cell central metabolism, which were simply grouped into six functional categories (Table 3): glycolysis and TCA cycle, pentose phosphate pathway, purine pathway and other PurR-regulated genes, riboflavin biosynthesis and transport, glycine biosynthesis, and nitrogen metabolism. The nucleotide sequences of six genes were identified to contain mutations in RH33, including ribC (the riboflavin kinase and FAD synthase gene), tkt (the transketolase

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Table 3 Relative expression changes and nucleotide sequence mutations of selected genes directly or indirectly participated in riboflavin biosynthesis in RH33. Gene product

Mutations

Glycolysis and TCA cycle pycA 1.32 pdhA 0.68 pdhB 0.72 pdhC 0.88 pdhD 0.88 gapB 1.08 citZ 1.96 citG 0.47 pta 0.77 ldh (lctE) 1.17 ackA 0.80 alsS 0.82 alsD 0.76 ptsG 1.07

Pyruvate carboxylase Pyruvate dehydrogenase (E1 alpha subunit) Pyruvate dehydrogenase (E1 beta subunit) Pyruvate dehydrogenase (dihydrolipoamide acetyltransferase E2 subunit) Pyruvate dehydrogenase/2-oxoglutarate dehydrogenase (dihydrolipoamide dehydrogenase E3 subunit) Glyceraldehyde-3-phosphate dehydrogenase Citrate synthase II (major) Fumarate hydratase Phosphotransacetylase L-lactate dehydrogenase Acetate kinase Alpha-acetolactate synthase Alpha-acetolactate decarboxylase PTS glucose-specific enzyme IICBA component

No No No No No No No No No Gly29Gly No No No No

Pentose phosphate pathway zwf 1.44 gntZ 0.29 yqjI 1.11 tkt 0.89 rpe 0.90 ywlF 0.90 prs 1.00

Glucose-6-phosphate 1-dehydrogenase 6-Phosphogluconate dehydrogenase Similar to 6-phosphogluconate dehydrogenase Transketolase Ribulose-5-phosphate 3-epimerase Ribose-5-phosphate isomerase B Phosphoribosylpyrophosphate synthetase

No No No 17delA Val4Asp No No

Nitrogen metabolism yweB (rocG) glnA gltC gltA gltB nasF nasE nasD nasC nasB narG narH narJ narI

4.82 0.81 0.82 0.46 0.37 1.62 1.59 1.34 1.31 1.81 6.48 13.59 16.35 19.27

Glutamate dehydrogenase Glutamine synthetase Transcriptional activator of the glutamate synthase operon Glutamate synthase (large subunit) (EC: 1.4.1.13) Glutamate synthase (small subunit) (EC: 1.4.1.13) Uroporphyrin-III C-methyltransferase Assimilatory nitrite reductase (subunit) Assimilatory nitrite reductase (subunit) Assimilatory nitrate reductase (catalytic subunit) Assimilatory nitrate reductase (electron transfer subunit) Nitrate reductase (alpha subunit) Nitrate reductase (beta subunit) Nitrate reductase (protein J) Nitrate reductase (gamma subunit)

No Glu65Lys No No No No No No No No No No No No

Glycine biosynthesis serA serC glyA folD

0.37 0.71 0.68 0.6

Phosphoglycerate dehydrogenase Phosphoserine aminotransferase Serine hydroxymethyltransferase Methylenetetrahydrofolate dehydrogenase/methenyltetrahydrofolate cyclohydrolase

No No No No

Gene

DFold

Purine biosynthesis pathway and other PurR-regulated genes purE 0.47 Phosphoribosylaminoimidazole carboxylase I (EC: 4.1.1.21) purK 0.48 Phosphoribosylaminoimidazole carboxylase II (EC: 4.1.1.21) purB 0.28 Adenylosuccinate lyase (EC: 4.3.2.2) purC 0.21 Phosphoribosylaminoimidazole succinocarboxamide synthetase (EC: 6.3.2.6) yexA (purS) 0.23 Required for phosphoribosylformylglycinamidine synthetase activity purL 0.24 Phosphoribosylformylglycinamidine synthetase II (EC: 6.3.5.3) purQ 0.24 Phosphoribosylformylglycinamidine synthetase I (EC: 6.3.5.3) Glutamine phosphoribosylpyrophosphate amidotransferase (EC: 2.4.2.14) purF 0.38 purM 0.30 Phosphoribosylaminoimidazole synthetase (EC: 6.3.3.1) purN 0.37 Phosphoribosylglycinamide formyltransferase (EC: 2.1.2.2) purH 0.50 Phosphoribosylaminoimidazole carboxy formyl formyltransferase/inosine-monophosphate cyclohydrolase purD 0.50 Phosphoribosylglycinamide synthetase purR 0.98 Transcriptional repressor of the purine operon purA 0.55 Adenylosuccinate synthetase guaB 0.88 Inosine-monophosphate dehydrogenase guaC (yumD) 0.23 GMP reductase yebB (pbuG) 0.42 Hypoxanthine/guanine permease Xpt 0.36 Xanthine phosphoribosyltransferase pbuX 0.3 Xanthine permease yqhZ 0.74 Probable transcription termination ytiP (pbuO) 0.24 Unknown

No No No No No No No No No No No No No No No Leu305Ile No No No No No

Riboflavin biosynthesis and transport ribA 13.27 ribB 13.19 ribC 3.26 ribG 21.74

No No Gly199Asp No

ribH ribT (ribD) ypaA

10.62 7.13 4.70

GTP cyclohydrolase II and 3,4-dihydroxy-2-butanone 4-phosphate synthase (EC: 3.5.4.25) Riboflavin synthase alpha chain [EC: 2.5.1.9] Riboflavin kinase and FAD synthase (EC: 2.7.1.26 and EC: 2.7.7.2) Diaminohydroxyphosphoribosylaminopyrimidine deaminase/5-amino-6-(5-phosphoribosylamino)uracil reductase [EC: 3.5.4.26 1.1.1.193] Riboflavin synthase (beta subunit) (EC: 2.5.1.9) Reductase A possible transporter of riboflavin

No No No

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gene), guaC (GMP reductase), ldh (L-lactate dehydrogenase), glnA (the glutamine synthetase gene), and rpe (the ribulose-5phosphate 3-epimerase gene). The mutations in the ribC and tkt were considered as beneficial mutations for riboflavin biosynthesis as discussed below, while causal links to riboflavin production were not obvious for the other mutations except the synonymous mutation in gene ldh, Gly(GGG)-29-Gly(GGA).

3.3. Transcriptional regulation and nucleotide mutation contributed to the riboflavin overproducing traits in B. subtilis RH33 First, in the riboflavin biosynthesis metabolism, the transcription of the rib genes, which are organized in an operon and encode the riboflavin biosynthesis pathway in B. subtilis, were strongly up-regulated in RH33 (Table 3; Fig. 1). The expression of the rib operon genes were significantly up-regulated by 13.2 (ribA), 13.2 (ribB), 10.6 (ribH), and 21.7-fold (ribG), reflecting their overexpression and direct contribution to riboflavin overproducing phenotype. Since rib operon genes are directly involved in riboflavin biosynthesis (Perkins et al., 1999), it is likely that the up-regulation of rib operon genes positively contributed to the riboflavin production. In B. subtilis, the rib operon has an untranslated regulatory leader region of 300 base pairs in front of its first gene, ribG. The leader sequence, a conserved regulatory element called RFN element, is responsible for negative regulation of the operon by FMN binding, which prevents the expression of the downstream genes by transcriptional or translational mechanisms (Nudler and Mironov, 2004). Revealed by DNA sequencing, there are no mutations in the RFN element in RH33. The up-regulation of the bifunctional flavokinase/FAD synthase gene ribC (3.3-fold), which converts riboflavin to FMN and FAD, is not beneficial for high riboflavin accumulation. However, DNA sequencing indicated that ribC of RH33 had a Gly-199-Asp mutation, which was coincident with the report of Bresler et al. (1973) and could lead up to 95% drop in enzyme activity. Thus, the up-regulation of rib operon in RH33 may be partly attributed to diminished feedback inhibition of FMN resulting from the low flavokinase activity of the mutated ribC. In addition to the riboflavin biosynthesis genes, the RFN element was also observed in the upstream of ypaA (Nudler and Mironov, 2004) which was predicted to encode a transporter of riboflavin. In RH33, ypaA gene was up-regulated by 4.7-fold. Based on these data, it was likely that the FMN concentration was maintained at a low level, reflected from the highly up-regulation of rib operon, and ypaA gene that all have the RFN element (Lee et al., 2001; Vogl et al., 2007). The mutation in ribC, which leads up to 95% drop in enzyme activity (Bresler et al., 1973), may directly contribute to the low FMN pool. Second, transcriptome profiling revealed that the mRNA expression level of the genes involved in the central carbon metabolism had small variations between wild-type strain 168 and the riboflavin overproducing strain RH33 (Table 3; Fig. 1). These genes may be highly expressed in both strains, as qualitatively reflected from their high absolute fluorescence intensities in one-color DNA microarray hybridization. Though riboflavin was over-produced, few genes involved in central metabolism showed differential expression in strain RH33. It may indicate that precursor supply due to the carbon flux through central metabolism was sufficient in RH33 under riboflavin overproduction condition. Alteration of carbon flow into the pentose phosphate pathway will affect the riboflavin production because the pentose phosphate pathway is a major source for the supply of ribose (or ribulose-5-phosphate), which is the starting point of riboflavin biosynthesis (Sauer et al., 1996; Zhu et al., 2006). As shown in Fig.

1, glucose-6-phosphate enters the pentose phosphate pathway after being oxidated to gluconate-6-phosphate by glucose-6phosphate dehydrogenase (zwf) and then is subsequently oxidized to ribulose-5-phosphate by gluconate-6-phosphate dehydrogenase. It was found that the transcription of gntZ, which was considered encoding the major 6-P-gluconate dehydrogenase, was significantly down-regulate (0.29-fold) in B. subtilis RH33. However, according to a recent report (Zamboni et al., 2004b), its isoenzyme gene yqjI was shown to have the major 6-P-gluconate dehydrogenase activity in B. subtilis, which was moderately upregulated in RH33 (1.1-fold) to compensate the enzyme activity and carbon flux. It should be noted that the expression of the transketolase gene tkt did not alter significantly in magnitude (0.89-fold). However, there was a frame-shift mutation in its coding region. It had been shown that the B. subtilis RH33 grew very slowly in minimal medium supplement with xylose or arabinose as the sole carbon source (unpublished data), which may be related to a dramatic lose of the transketolase activity. Indeed, B. subtilis RH33 showed only 4% transketolase activity compared with that of B. subtilis 168 (177 nmol min1 mg1 protein). The low activity of transketolase may decrease the shunt of pentose back into glycolysis, allowing cells to redirect more carbon flux through the oxidative pentose pathway to the purine nucleotide biosynthesis pathway, which provided the precursors of riboflavin production (Kamada et al., 2001). In short, tkt contributed to riboflavin overproduction trait of RH33, not at the level of transcriptional regulation, but through frame-shift mutation to reduce enzyme activity of gene tkt.

3.4. Identification of metabolic bottleneck for increasing riboflavin production in B. subtilis RH33 Purine nucleotides, involving in biosynthesis of riboflavin by providing the immediate precursor GTP, are synthesized by purine genes (purEKBCSQLFMNHD). Unexpectedly, the purine genes together with glycine biosynthesis genes (serA, serC, glyA, folD) (Saxild et al., 2001) were all greatly down-regulated in RH33 (Table 3; Fig. 1). The down-regulation of purF, purD, glyA, and folD were also revealed by quantitative RT-PCR experiment (Table 2). Thus, the down-regulated purine and glycine (another building block for riboflavin) biosynthesis genes would limit the supply of the precursors for riboflavin overproduction (Heefner et al., 1988; Jimenez et al., 2005; Mateos et al., 2006; Schlupen et al., 2003; Stahmann et al., 2000). Nonetheless, it was interesting to note that the downregulated purine, glycine, and other genes involved in purine metabolism (guaC, pbuG, xpt-pbuX, yqh, and pbuO) were all negatively regulated by global regulator PurR in B. subtilis (Saxild et al., 2001). The nucleotide sequencing results indicated that there were no mutations in the regulatory region (PRPP binding motif) and coding region of the purR gene. Among all the PurRregulated genes only guaC had a point mutation (Table 3). It was reported that the defective GMP reductase (guaC) that reduces conversion of GMP to IMP had been identified in a B. subtilis riboflavin overproducer (Nygaard, 1993). However, the mutation of guaC in B. subtilis RH33 was neutral (from Leu to Ile) and probably had no effect on protein function. Since no mutations were found in the regulatory region and coding region of the PurR-regulated genes (including purR) except guaC, the down-regulation of those genes may be due to the repression by PurR. It binds to the operator sites (PurBox) of its target genes and inhibits their transcription. According to the current model, the PRPP can bind PurR to prevent its binding to DNA and lead to derepression of PurR-regulated genes (Weng et al., 1995). Therefore, it can be deduced that the low expression

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of PurR-regulated genes might be caused by a small cellular PRPP pool. Subsequently, metabolite analysis was carried out and tested this hypothesis (Table 4). The PRPP pool in B. subtilis RH33 was only 24% that of B. subtilis 168 measured at the conditions as the microarray experiments. In addition, PRPP itself is an important metabolite precursor for purine biosynthesis as it is required in the de novo and salvage pathways of purine biosynthesis (Jimenez et al., 2008). It was proposed that enhancing PRPP pool would derepress the expression of pur operon and glycine biosynthesis genes, thus increasing the precursors supply and leading to increased production of riboflavin in B. subtilis RH33. At present, there is only one report employing the strategy of modulating the PRPP pool for riboflavin production (Jimenez et al., 2008) in a hypothesis-driven approach.

249

The selected prs gene was overexpressed in B. subtilis RH33 under the control of the P43 promoter to construct the strain B. subtilis RH33-Prs. The enzyme activities of PRPP synthetase were 3.070.7 and 10.172.0 nmol min1 mg1 protein for B. subtilis RH33 and B. subtilis RH33-Prs, respectively, indicating that PRPP synthetase was overexpressed successfully. To assess the effects of the manipulation on the physiological parameters, batch cultivations were carried out in minimal medium. The results displayed only minor variability in the specific cell growth rate, specific glucose uptake rate, specific organic acids secretion rate, and specific riboflavin production rate between B. subtilis RH33 and B. subtilis RH33-Prs (Table 6). Unexpectedly, the PRPP concentration was almost constant in the mutant (Table 4). These indicated that overexpression of prs gene alone had little effect on the supply of PRPP and on specific riboflavin production rate.

3.5. Effect of prs overexpression on riboflavin production PRPP is an important metabolite required for purine, pyrimidine, tryptophan, and histidine biosynthesis. As discussed above, the low concentration of PRPP may be the bottleneck for further increasing riboflavin production in B. subtilis RH33. The formation of PRPP is catalyzed by the enzyme PRPP synthetase, which is encoded by prs gene. The expression level of prs was not notably altered. In addition, the enzyme activity assay showed that the enzyme activity of RPPP synthetase was almost the same between B. subtilis RH33 and B. subtilis 168 (Table 5). Therefore, it was reasoned that amplification of prs gene was expected to increase the supply of PRPP and consequently result in increased riboflavin production.

3.6. Enhancement of riboflavin production by co-expression of prs and ywlF genes As the strain B. subtilis RH33-Prs did not show the expected traits in improving PRPP supply, to diagnose the problem(s), further measurements of precursor concentrations of the metabolites were carried out to elucidate the limitations. Compared to RH33, ribulose-5-phosphate had similar levels but ribose5-phosphate had significantly lower concentrations in RH33Prs (Table 4). The insufficient ribose-5-phosphate pool was identified as the limiting factor for the indistinctive effect on increasing RPPP concentration and riboflavin production in B. subtilis RH33-Prs.

Table 4 Intracellular metabolite concentration in cell extracts of B. subtilis. Metabolite [nmol mg1 (dry wt)]

Ribose-5-P Ribulose-5-P

b

RH33 a

PRPP

a

168

RH33-Prs a

2.5070.36 3.6270.45b 1.1170.29a 4.3071.54a

0.8870.31 0.8770.38b 1.2870.33a 4.3671.52a

RH33-PY a

0.9170.32 0.9870.39b 0.5570.25a 4.1871.46a

4.1470.60a 5.7770.65b 1.9870.40a 4.2071.52a

Sampled during exponential growth period cultivated in minimal medium. Sampled during exponential growth period cultivated in LBG medium.

Table 5 Specific enzyme activities in cell extracts of B. subtilis, sampled during exponential growth period. Enzyme activities (nmol min1 mg1 protein)

168

RH33

RH33-Prs

RH33-PY

PRPP synthetase Ribose-5-phosphate isomerase B

4.170.8 28.772.5

3.070.7 22.372.5

10.172.0 27.472.5

10.772.0 61.475.5

Table 6 Metabolic characterization of B. subtilis strains during exponential growth period (OD600 0.3–0.6) cultivated in minimal medium. Parameter

168

Specific glucose uptake rate (mmol g1 CDW h1) Specific growth rate (h1) Specific organic acidsa secretion rate (mmol g1 CDW h1)

4.570.4 0.3170.03

4.270.4 0.2770.02

4.170.4 0.2870.03

4.270.4 0.2770.03

1.6370.31 NDb

1.5170.28 0.04570.002

1.4770.22 0.04770.003

1.3070.22 0.05370.003

Specific riboflavin production rate (mmol g1 CDW h1)

RH33

Results represent the mean values with standard deviations from three independent measurements. a Organic acid including acetate, lactate, and pyruvate. b Not detected.

RH33-Prs

RH33-PY

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Relative gene expression (RH33-RY/RH33)

7 6 5 4 3

10 9

Glucose (g/L)

8 7 6 5 4 3 2 1 0 10 9 8 Biomass (0D600)

As shown in Fig. 1, ribose-5-phosphate can be generated from ribulose-5-phosphate catalyzed by ywlF encoding ribose-5-phosphate isomerase B. We decided to co-overexpress the prs and ywlF genes simultaneously. In this study, the plasmid containing both prs gene and ywlF gene under the control of the P43 promoter was introduced into the prs locus of the B. subtilis RH33 chromosome to construct B. subtilis RH33-PY strain. Both PRPP synthetase and ribose-5-phosphate isomerase B enzyme activities were significantly higher than strain RH33, as shown in Table 5, indicating the successful overexpression of prs and ywlF genes. When both prs and ywlF were overexpressed, the specific cell growth rate and glucose uptake rate were almost the same compared with RH33 (Table 6). Meanwhile, RH33-PY had a slightly lower organic acid secretion. The productivity of riboflavin was significantly increased from 0.04570.002 to 0.0537 0.003 mmol g1 CDW h1. The concentration of intermediate metabolites PRPP and ribose-5-phosphate were much higher in B. subtilis RH33-PY compared to RH33 strain (Table 4), which implied that the higher yield of riboflavin was attributable to the sufficient supply of PRPP. Although the overexpression of ribose-5-phosphate isomerase B caused more carbon flux to redirect from ribulose-5-phosphate to ribose-5-phosphate, the concentrations of ribulose-5-phosphate in these strains were almost constant, which indicated the abundant precursor supply provided by central carbon metabolism in B. subtilis RH33 as mentioned above. PRPP has dual roles in riboflavin production: one is used as a precursor, and the other is to bind PurR and depress PurRregulated genes. To confirm that indeed the increased PRPP pool resulted in up-regulation of PurR controlled genes in RH33-PY strain, a quantitative RT-PCR experiment was conducted to measure the transcript abundances of PurR-regulated genes relative to parental strain RH33. The fold change of the selected PurR-regulated genes, including purE, purD, guaC, purA, glyA, folD, pbuG, xpt, pbuX, ytiP, and yqhZ, were shown in Fig. 2. As expected, expression of those PurR-regulated genes, including the purine pathway genes (purE, purD) and the glycine biosynthesis pathway genes (glyA, folD), were induced in RH33-PY (Fig. 2), which was in accordance with our hypothesis. In B. subtilis RH33-PY, it is suggested that the derepressed PurR-regulated genes activated the pathways that supplying precursors for riboflavin biosynthesis, which in turn led to the increase of riboflavin production.

7 6 5 4 3 2 1 0 250

Riboflavin Concentration (mg/L)

250

200 150 100 50 0 0

2

4

6

8

10

Time (h) Fig. 3. Time-course profiles of biomass, glucose, and riboflavin concentration of B. subtilis RH33 (closed triangles), RH33-Prs (closed squares) and RH33-PY (closed circles) in LBG medium during shake flask batch cultivations.

Fig. 3 shows the time-course profiles of riboflavin production, biomass, and glucose concentration exhibited by RH33, RH33Prs, and RH33-PY grown in shake flask with LBG medium. The cell growth and glucose uptake profiles of RH33-Prs and RH33PY were almost the same as the parental strain RH33. Little effect on riboflavin production was observed in RH33-Prs compared with RH33. However, the RH33-PY mutant exhibited a riboflavin titer of 3273% higher than RH33 (Fig. 3). Thus, it indicated that the metabolic engineering strategy identified by transcriptome analysis successfully improved the trait of riboflavin overproduction in RH33 on a shake flask scale. 3.7. Riboflavin production in fed-batch fermentation

2 1 0 pur E pur D pur A gly A fol D guaC pbu G xpt

pbuX ytiP yqhZ

Genes Fig. 2. The fold change of selected PurR-regulated transcripts in B. subtilis RH33PY, relative to RH33. Values represent mean and s.d. (n ¼ 3).

To investigate the potential of the strain B. subtilis RH33-PY in a larger scale fermentor, we tested the performance of strain B. subtilis RH33-PY in fed-batch fermentation using the feeding profile as described in Materials and methods. The cell growth rates were also very similar among the parental strain RH33 and two engineered strains RH33-Prs and RH33-PY (Fig. 4A). Compared to parental strain RH33, an average of 25% increase in riboflavin titer, up to 15 g l1, was achieved in RH33-PY (Fig. 4B), while the improvement was only about 6% in strain RH33-Prs

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140

Biomass (OD600)

120 100 80 60 40 20 0 0

6

12

18 24 30 Feed time (h)

36

42

48

18 Riboflavin concentration (g/L)

16 14

251

analysis. By co-overexpressing prs and ywlF in RH33, PRPP pool level was significantly increased. This had dual effects: one was to derepress the purine genes negatively regulated by PurR (Fig. 2), and the other was to provide more precursors for purine biosynthesis pathways. The strategy developed in this work allowed for a 25% increase in riboflavin titer, up to 15 g l1, compared to the parental strain during a 5-l fed-batch fermentation. An increasing number of publications have demonstrated how transcriptome analysis has been successfully applied to understand and engineering metabolism of the organisms. This trend will be expected to continue in the post-genomic era. Unfortunately, elucidating and application of the mechanism(s) underlying the improved performance of the strains developed by random mutagenesis and selection are still limited, partly due to the complexities of metabolic and regulatory networks encoded in the even simplest bacterial genome. The whole-genome gene expression analysis, integrated with nucleotide sequencing and metabolite measurements, represents a powerful approach to exploring and exploiting of genomic data, thus greatly facilitating the identification of novel targets for strain improvement. This systems strategy should be in principle applied for rational microbial development by deducing the global physiology and regulation of an overproducer strain that is obtained by combinatorial approaches.

12 Acknowledgments

10 8 6 4 2 0 0

6

12

18 24 30 Feed time (h)

36

42

48

Fig. 4. Time-course profiles of biomass (A) and riboflavin concentration (B) of B. subtilis RH33 (closed triangles), RH33-Prs (closed squares) and RH33-PY (closed circles) during fed-batch cultivations in a 5-l bioreactor.

This research was supported by the National Natural Science Foundation of China (NSFC-20536040), the National Project of Key Fundamental Research (2007CB707802), the Development Project of Science and Technology of Tianjin (05YFGZGX04500) and Programme of Introducing Talents of Discipline to Universities (B06006). CapitalBio Co. Ltd., in Beijing was acknowledged for help with the microarray experiment. The microarray data described in this publication had been deposited in the public database (Gene Expression Omnibus, GEO) with accession number of GSE12873.

Appendix A. Supplementary material 1

(12.7 g l ). Therefore, in fed-batch fermentation, modulating PRPP pool by co-overexpressing prs and ywlF successfully improved riboflavin production in B. subtilis while causing little effect on cell growth. These data also suggested that increasing PRPP pool may be a general strategy to improve riboflavin production. The alterations of riboflavin production in the three strains have the same trend in the medium tested. Another successful example for enhancing riboflavin production by manipulation of the PRPP pool was achieved in Ashbya gossypii (Jimenez et al., 2008). In this study, based on known regulatory and metabolic information and new insights generated from transcriptome analysis, combined with nucleotide sequencing and intracellular metabolite concentration measurement, metabolic bottleneck and novel target genes were identified and co-amplified to further enhance production of riboflavin in B. subtilis. Microarray analysis identified that purine and glycine biosynthesis genes were significantly down-regulated in the riboflavin-producing strain RH-33. These genes were repressed by the global regulator PurR whose activity was modulated by small molecular PRPP, which is also the precursor for purine synthesis. It was inferred that the reduced expression of PurR-regulated genes might be caused by a low PRPP pool, which was subsequently confirmed by metabolite

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.ymben.2009.05.002.

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