Analysis of the Transcriptional Regulator GlpR, Promoter Elements,and Posttranscriptional Processing Involved in Fructose-InducedActivation of the Phosphoenolpyruvate-Dependent SugarPhosphotransferase System in Haloferax mediterranei

Lei Cai,a,b Shuangfeng Cai,a,b Dahe Zhao,a Jinhua Wu,a,b Lei Wang,a Xiaoqing Liu,a,b Ming Li,a,b Jing Hou,a,b Jian Zhou,a Jingfang Liu,a

Jing Han,a Hua Xianga

State Key Laboratory of Microbial Resources, Institute of Microbiology, Chinese Academy of Sciences, Beijing, Chinaa; University of Chinese Academy of Sciences, Beijing,Chinab

Among all known archaeal strains, the phosphoenolpyruvate-dependent phosphotransferase system (PTS) for fructose utiliza-tion is used primarily by haloarchaea, which thrive in hypersaline environments, whereas the molecular details of the regulationof the archaeal PTS under fructose induction remain unclear. In this study, we present a comprehensive examination of the regu-latory mechanism of the fructose PTS in the haloarchaeon Haloferax mediterranei. With gene knockout and complementation,microarray analysis, and chromatin immunoprecipitation-quantitative PCR (ChIP-qPCR), we revealed that GlpR is the indis-pensable activator, which specifically binds to the PTS promoter (PPTS) during fructose induction. Further promoter-scanningmutation indicated that three sites located upstream of the H. mediterranei PPTS, which are conserved in most haloarchaealPPTSs, are involved in this induction. Interestingly, two PTS transcripts (named T8 and T17) with different lengths of 5= untrans-lated region (UTR) were observed, and promoter or 5=UTR swap experiments indicated that the shorter 5=UTR was most likelygenerated from the longer one. Notably, the translation efficiency of the transcript with this shorter 5=UTR was significantlyhigher and the ratio of T8 (with the shorter 5=UTR) to T17 increased during fructose induction, implying that a posttranscrip-tional mechanism is also involved in PTS activation. With these insights into the molecular regulation of the haloarchaeal PTS,we have proposed a working model for haloarchaea in response to environmental fructose.

The phosphoenolpyruvate (PEP)-dependent sugar phospho-transferase system (PTS) uses PEP as the phosphoryl donor to

phosphorylate sugars for transport into cells (1, 2). A typical PTScontains five proteins, PtsI (or EI), HPr, PtsA, PtsB, and PtsC.Phosphotransfer from PEP to sugar is mediated by these five pro-teins in a cascade, and the PtsC component at the end of thecascade couples phosphorylation with the translocation of thespecific sugars (3, 4). The PTS is an important apparatus for sugaruptake and degradation in bacteria, and most bacteria have beenshown to possess at least one complete PTS (4). The PTSs of bac-teria can sense the primary metabolic or environmental signal andturn on the uptake system (5, 6). In response to the environmentalsignal, the derivatives of sugars always serve as positive or negativeeffectors, while the global or specific transcriptional regulators, incooperation with cyclic AMP (cAMP) or primary metabolites, areinvolved in the complicated regulation of the PTS via direct bind-ing to the promoter regions of PTS genes (2, 7).

In contrast to the case for bacteria, research on archaeal PTSshas received attention just in the past few years. The first reportthat archaea have PTS genes was published in 2006, based on thegenome sequencing of the haloarchaeon Haloquadratum walsbyi(8). Recent studies of haloarchaeal genome sequences have indi-cated that many haloarchaea contain PTS genes (8–11), and 6 outof 24 haloarchaeal genomes have a complete fructose-specific PTSgene cluster, including Haloterrigena turkmenica, Halalkalicoccusjeotgali, Haloarcula hispanica, Haloarcula marismortui, Haloferaxvolcanii, and Haloferax mediterranei (11). Recently, a functionalfructose-specific PTS has been identified in H. volcanii using ge-netic methods, and fructose was shown to be able to upregulate

the transcription of this PTS gene cluster (12), but the moleculardetails of the fructose-induced PTS activation in archaea remainunclear. Interestingly, earlier research on H. volcanii indicates thata DeoR family transcriptional regulator, GlpR, represses the ex-pression of fructose and glucose metabolic enzymes (2-keto-3-deoxy-D-gluconate kinase [KDGK] and phosphofructokinase[PFK]) at the transcriptional level when cells are grown on glyc-erol (13). In addition, it was reported that glpR is cotranscribedwith the downstream phosphofructokinase gene (fruK) (13). ThePTS gene cluster is located just adjacent to glpR-fruK. As an im-portant regulator which usually functions in sugar metabolism inbacteria (14, 15), GlpR is probably involved in the transcriptionalregulation of the haloarchaeal PTS cluster. However, as far as weknow, the relationship between GlpR and the PTS activation inhaloarchaea has not yet been established.

Recently, the genome of H. mediterranei was completely se-quenced by our laboratory (16). A genome-wide in silico analysisshowed that the arrangement of fructose metabolism-relatedgenes, including those of the PTS system in H. mediterranei, is

Received 10 October 2013 Accepted 10 December 2013

Published ahead of print 13 December 2013

Address correspondence to Hua Xiang, xiangh@im.ac.cn.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.03372-13.

Copyright © 2014, American Society for Microbiology. All Rights Reserved.

doi:10.1128/AEM.03372-13

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identical to that of H. volcanii. However, in contrast to H. volcanii,H. mediterranei can synthesize biodegradable polymers such aspolyhydroxyalkanoates (PHA) from many inexpensive carbon re-sources (17–19). Studying the mechanism of regulation of the PTSin H. mediterranei not only is useful to compare the different generegulation strategies between bacteria and archaea but can alsoresult in a deeper understanding of the carbon sensing and utili-zation by this specific haloarchaeal PHA producer. In the presentstudy, a comprehensive investigation of the regulatory mecha-nism of the PTS was performed in H. mediterranei. We demon-strated that GlpR is an indispensable activator of the PTS genecluster upon fructose induction via direct binding to the PTS pro-moter region (PPTS). Interestingly, we also revealed an additionalposttranscriptional mechanism which could increase the transla-tion efficiency of PTS transcripts. Together, our results help elu-cidate the complex and delicate mechanisms of fructose PTS reg-ulation in the domain of archaea.

MATERIALS AND METHODSStrains and growth conditions. The strains used in this study are listed inTable S1 in the supplemental material. Escherichia coli JM109 was used asthe host for the cloning experiments (Novagen, Madison, WI, USA) andwas grown in Luria-Bertani medium at 37°C (20). Unless otherwise noted,H. mediterranei DF50 (21) and the gene knockout mutants were culti-vated at 37°C in nutrient-rich AS-168L medium (22), and H. mediterraneistrains harboring expression plasmids were cultivated in AS-168SYL me-dium (AS-168L without yeast extract) (22). Chemically defined medium(CDM) [consisting of (per liter) 150 g NaCl, 20 g MgSO4 · 7H2O, 2 g KCl,50 mg FeSO4 · 7H2O, 0.36 mg MnCl2 · 4H2O, 5 g NH4Cl, and 15 g piper-azine-N,N=-bis(2-ethanesulfonic acid) PIPES, pH 7.2] with different con-centrations of fructose or glucose was used to verify the utilization of thecarbon source by H. mediterranei mutant strains. When required, ampi-cillin, uracil, and 5-fluoroorotic acid (5-FOA) were added to the media atfinal concentrations of 100 mg/liter, 50 mg/liter, and 250 mg/liter, respec-tively.

Gene knockout and complementation. In-frame deletion and com-plementation strains were generated according to previously publishedprotocols (21, 23). All the primers used in this study are listed in Table S2in the supplemental material, and the plasmids are listed in Table S1 in thesupplemental material. The transformation of H. mediterranei was per-formed by the polyethylene glycol-mediated spheroplast transformationmethod (24). The plasmid sequences and mutant strains were verified byPCR and DNA sequencing.

RNA extraction, quantitative reverse transcription-PCR (qRT-PCR), and circularized RNA (CR)-RT-PCR. H. mediterranei DF50 cellsand the gene knockout mutants were cultured at 37°C in AS-168L me-dium. When the optical density at 600 nm (OD600) reached 1.5, glucose orfructose was added to the medium to a final concentration of 50 mM, andthe cells were incubated for 45 min. The sugar-induced cells (3 ml) werethen immediately collected for RNA extraction using TRIzol reagent (In-vitrogen, Carlsbad, CA, USA) according to the manufacturer’s instruc-tions. An equal volume of TBSL buffer (consisting of [per liter] 150 gNaCl, 20 g KCl, 5 g MgSO4 · 7H2O, and 100 mM Tris-HCl, pH 7.1) (22)was added to the cells in the control group. To remove DNA contamina-tion, DNase I (Promega, Madison, WI, USA) digestion was performed on1 �g of diluted RNA.

The specific primer pairs for the target DNA regions (see Table S2 inthe supplemental material) and suitable concentrations of the cDNA tem-plates or genomic DNA were used for quantitative PCR (qPCR). Theamplification and detection of target regions were performed on a Rotor-Gene Q real-time cycler (Qiagen, Valencia, CA) under a standard three-step PCR procedure (including initial denaturation at 95°C for 10 minfollowed by 40 cycles of denaturation at 95°C for 30 s, annealing at 55°Cfor 30 s and synthesis at 72°C for 30 s; a melting curve was generated by

linear heating from 70°C to 95°C over 25 min). For the synthesis of thecDNA, 200 ng of DNase I-treated total RNA was reverse transcribed viarandom hexamer primers by using the Moloney murine leukemia virusreverse transcriptase (M-MLV-RT) (Promega, Madison, WI, USA).DNase I-treated RNA (without reverse transcription) was used to checkfor genomic DNA contamination.

CR-RT-PCR (25, 26) was used to determine the 5= untranslated region(UTR) of the PTS gene cluster. RNA circularization was carried out asdescribed previously (25). Self-ligated RNA was reverse transcribed viarandom hexamers primers as described above. The cDNA was first am-plified with a gene-specific primer pair, cRT1F and cRT1R, and a secondPCR was performed to enhance the specificity by using an inner primerpair, cRT2F and cRT2R. The products of the second PCR were cloned intothe TA vector pUCm-T (Sangon Biotech, Shanghai, China) according tostandard procedures, and 15 clones from each RNA sample were analyzedby sequencing.

Constructs used for transformation of H. mediterranei. For analysisof the promoter activity in vivo, a plasmid-based transcriptional reportersystem using a soluble modified red-shifted green fluorescent protein(smRSGFP) (27) was constructed as previously described (28). All theplasmids used to transform H. mediterranei cells were derived frompWL502, and the details of their constructions are shown in the supple-mental material. The plasmids pL117, pPR, and pPF were used to analyzethe wild-type promoter activities of PTS (PPTS), glpR (PglpR) and fba (Pfba),respectively. To analyze the regulatory region of PPTS, 5= flanking deletionmutants of PPTS (named pL93, pL77, pL56, pL24, and pGFP-0) and scan-ning site-directed mutants (from pM�1216 to pM7975) were trans-formed into H. mediterranei DF50 to detect the mutated promoter activ-ity. The activities of GlpR and Myc-tagged GlpR were determined byintroducing the plasmids pL117CR and pL117Rm, respectively, into theH. mediterranei DF50 �glpR strain. In the promoter or 5= UTR swapexperiment, pHSP, pUTR-M, pT8, and pT17 were generated by con-structing fusions of the promoter and 5=UTR regions between hsp5 (am-plified from the plasmid pSCM307 [29]) and the PTS gene cluster.

Analysis of the smRSGFP fusion reporter system. The fluorescenceintensity of each smRSGFP fusion reporter plasmid-harboring strain wasmeasured using a Synergy H4 hybrid microplate reader (BioTek Instru-ments Inc., Winooski, VT, USA). The excitation wavelength was set to 488nm, and the emission wavelength was 509 nm (27). All strains were incu-bated at 37°C in AS-168SYL medium until the OD600 reached 1.5; 90 �l ofeach culture was then transferred to black polystyrene 96-well plates(3916; Costar, Corning, NY, USA). In most cases, 10 �l of a fructose orglucose stock solution was added to the designated induction wells on the96-well plates to a final concentration of 50 mM. For tests of inductionwith the metabolic intermediate fructose-1-phosphate (F-1-P), the finalconcentrations of F-1-P and fructose were reduced to 2 mM. In each assay,10 �l of TBSL buffer was used as a negative control for basal fluorescenceintensity. All of the plates were incubated at 37°C for 8 h before measure-ment.

Primer extension. H. mediterranei strains harboring different plas-mids were used in the primer extension assay with a specific primer basedon the sequence of the GFP gene. Primer extension reactions were per-formed using 5 �g of total RNA and 3 pmol of the 5=-biotin-labeledprimer gfpRbio (see Table S2 in the supplemental material) with the re-verse transcription protocol described above. The extension productswere analyzed on an 8% acrylamide sequencing gel. A chemiluminescentnucleic acid detection module kit (Pierce-Thermo Scientific, Rockford,IL) was used for biotin detection.

Immunoprecipitation. The interaction between GlpR and the pro-moter region was analyzed by the chromatin immunoprecipitation(ChIP) assay. Myc-tagged GlpR was expressed using the pRm plasmid (seeTable S1 in the supplemental material) in the H. mediterranei �glpRstrain. Cells were harvested at the mid-logarithmic phase (OD600 � 1.5)with or without fructose induction. The ChIP experiments were per-formed according to a previously described protocol (30, 31). The enrich-

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ment of genomic fragments was analyzed by qPCR with the input DNAsamples as controls. The primers used are listed in Table S2 in the supple-mental material. The PCR and thermocycling conditions were the same asdescribed above for qRT-PCR. Each ChIP assay had five biological repli-cates. In each ChIP sample, the enrichment of GlpR-Myc interacting witheach locus was calculated compared to the input sample using relativequantitation.

Microarray assay and deep sequencing. The RNA samples used forqRT-PCR from DF50 cells with or without fructose induction were sub-jected to microarray analysis. Oligonucleotide microarrays were designedand manufactured by Capital Bio and Agilent Technologies based on thewhole genomic sequence of H. mediterranei. The microarray assay wascarried out as previously described (32). Each assay was repeated threetimes. The resulting data were analyzed by Significance Analysis of Mi-croarrays (SAM) software version 2.23b (33). The same protocols used forthe microarray assays were also performed on the total RNA samples fromthe �glpR mutant strain with or without fructose induction. The deepsequencing of the transcriptome of H. mediterranei was performed onHiSeq sequencing systems (Illumina HiSeq 2000) at the Beijing Instituteof Genomics of the Chinese Academy of Sciences.

Prediction of RNA secondary structures. The program Sfold (34, 35)was used for the prediction of putative secondary structures of RNA (http://sfold.wadsworth.org).

Microarray data accession number. The microarray data have beendeposited in the NCBI GEO library under accession number GSE41134.

RESULTSGlpR is essential for the activation of the PTS promoter by fruc-tose. In H. mediterranei, the PTS genes (HFX_1559 to HFX_1563)were organized in an operon corresponding to a polycistronictranscript (Fig. 1A). The functional involvement of the PTS infructose utilization was confirmed via genetic methods that wereused in H. volcanii (21) (see Fig. S1 and S2 in the supplementalmaterial). The DNA sequence of the intergenic region betweenfruK and ptsC is given in Fig. 1B. Deep sequencing of the total RNAof H. mediterranei DF50 indicated that there were two transcripts,with 17- and 8-nucleotide (nt) 5= untranslated regions (UTRs),respectively. The start site (A) (�1) of the longer transcript (T17)

is 9 nt away from the start site (G) (�10) of the shorter transcript(T8) (Fig. 1B and C), and these start sites were confirmed by CR-RT-PCR. However, only one typical promoter containing a puta-tive TFIIB recognition element (BRE) (�36 GAAAGG �31) and aputative TATA box (�30 ATTTTT �25) was found (Fig. 1B).

In silico analysis showed that the DeoR family transcriptionalregulator, GlpR, is highly conserved in haloarchaea (�60% iden-tity), but it is more distantly related to its homologs in other ar-chaea and bacteria (�45% identity) in comparison. The proteinsequence multiple-alignment analysis by BLASTP indicated thatapproximately 70 of the 255 amino acids in the N terminus ofGlpR form a putative helix-turn-helix (HTH) motif, and the re-maining amino acids near the C terminus form a DeoR-type reg-ulator C-terminal sensor domain. To determine whether the tran-scription of the PTS gene cluster is regulated by GlpR, H.mediterranei DF50 and the �glpR strain were analyzed by microar-ray assays with or without fructose induction (GEO accessionnumber GSE41134). The transcriptional fold changes of the PTSgene cluster and neighboring genes (HFX_1558 to -1565) arelisted in Table 1. It was shown that the PTS gene cluster and theglpR-fruK operon, which were highly upregulated by fructose inH. mediterranei DF50, were not inducible in the �glpR strain. Inaddition, the transcription of fba was not significantly changed ineither group (Table 1). These results strongly suggested that GlpRis an indispensable regulator in fructose-induced PTS activationin H. mediterranei.

To confirm the regulation of PTS by GlpR in vivo and to con-veniently investigate the PTS promoter, the �glpR strain harbor-ing different GFP-based reporter plasmids was investigated; theDF50 strain served as a positive control. It was shown that whetherinduced by fructose or not, the fluorescence intensity of the �glpRstrain harboring pL117, which expressed smRSGFP under controlof the wild-type PTS promoter (134 bp upstream of translationstart codon of ptsC), was much closer to the basal fluorescenceintensity of DF50 harboring pL117 (without fructose induction)

FIG 1 Map of the PTS gene cluster, the promoter sequence, and analysis of PTS transcripts. (A) Genetic organization of the H. mediterranei PTS gene cluster andneighboring genes. glpR (HFX_1565) encodes a protein homologous to a DeoR family transcriptional regulator, and fruK (HFX_1564) encodes the 1-phospho-fructokinase. ptsC, ptsA, hpr, ptsI, and ptsB (HFX_1563 to HFX_1559) encode a complete PTS, and fba (HFX_1558) encodes a fructose-1,6-bisphosphatealdolase. The primers used for CR-RT-PCR are indicated with arrows. (B) Promoter sequence of the PTS gene cluster is shown. The stop codon TAA of theupstream gene fruK and the start codon ATG of ptsC are boxed. The start sites (indicated by arrows) of the two transcripts containing 5=UTRs of different lengthswere determined by CR-RT-PCR. The putative TATA box and BRE are indicated by single and double underlines, respectively. (C) Statistical results of mRNAdeep sequencing. The counts (y axis) of corresponding nucleotides (x axis) of a 24-bp sequence of the PPTS are presented. The positions with the most significantincreases in sequencing counts are marked by arrows, which indicate the two start sites of PTS transcripts.

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(Fig. 2). The �glpR strain harboring pL117CR showed that theactivation of the PPTS by fructose was restored through the expres-sion of GlpR (using its native promoter, PglpR) in the �glpR strain(Fig. 2). Furthermore, the transcriptional activity of PglpR was alsoanalyzed via the reporter plasmid pPR, which expresses smRSGFPusing PglpR. The fluorescence intensity of the PglpR fusion reportersystem increased slightly (approximately 1.5-fold) when DF50cells were induced by fructose but did not change when �glpR cellswere tested (Fig. 2). In contrast, regardless of fructose induc-tion, the fluorescence intensity was similar in DF50(pPF) and�glpR(pPF) transformants (both expressing smRSGFP with thePfba promoter) (Fig. 2). These results confirmed that GlpR is es-sential for the fructose-induced transcriptional activation of thePTS and the glpR-fruK gene clusters, and it may act as a positiveregulator for fructose-induced PTS expression. This finding isquite interesting, as GlpR has been previously identified as a globalrepressor that inhibits the activities of KDGK and PFK in H. vol-canii when cells are cultured in a glycerol-based medium (13). Thedifferent functions of GlpR are likely attributable to the different

carbon sources (fructose versus glycerol) being used and/or todifferent promoters.

GlpR binds directly to the PPTS during fructose induction. Todetermine whether the activation of gene expression by GlpR oc-curs via direct binding to the promoter sequence, a ChIP assaycoupled with qPCR analysis was performed on the �glpR strainharboring pRm (a Myc-tagged GlpR expression plasmid) with orwithout fructose induction. The recombinant GlpR-Myc (ex-pressed by pL117Rm) was revealed to be able to restore the activityof wild-type GlpR in the �glpR strain (data not shown). ThreeDNA loci (FPTS, Fiic, and FphaE) were investigated for their inter-action with GlpR, with the F16S locus (a fragment of the 16S rRNAgene) used as an internal control for data normalization. FPTS (119bp) represented a fragment of the PPTS region (bp �92 to �27upstream of the PTS gene cluster), and Fiic (125 bp) representedthe intragenic region of ptsC located approximately 550 bp to 650bp downstream of FPTS. The FphaE locus (189 bp) containing thepromoter region of phaE (which encodes a subunit of polyhy-droxyalkanoate synthase in H. mediterranei [19]) was tested as anegative control because the expression level of phaE did notchange in the microarray experiment when the cells were treatedwith fructose (data not shown). After the induction by fructose,the FPTS locus exhibited a 2-fold enrichment of binding to GlpR-Myc over the negative-control locus FphaE, and the enrichments ofthe FPTS and FphaE loci were similar to each other in the absenceof fructose induction (Fig. 3). As expected, the fold enrichment ofthe Fiic locus remained unchanged and was similar to that of theFphaE locus, with or without fructose induction (Fig. 3). Theseresults demonstrated that GlpR could directly bind to PPTS whenH. mediterranei cells were treated with fructose, but the interac-tion between GlpR and PPTS was not apparent without fructoseinduction. The significantly increased binding between GlpR andPPTS under fructose induction indicated again that GlpR is anactivator of PTS transcription.

Three regions within PPTS account for fructose induction.The above results demonstrated that GlpR acts as a positive regu-lator for the induction of the PTS gene cluster by fructose viadirect binding to the PPTS. To experimentally analyze the cis-act-ing elements of the PPTS, 5= flanking deletion and site-directedmutagenesis analysis of the PPTS were carried out based on thesmRSGFP fusion reporter system of plasmid pL117. The fluores-cence intensities of H. mediterranei DF50 transformants harbor-ing deletions or site-directed mutagenesis constructs were mea-sured with or without fructose induction (Fig. 4).

The wild-type PTS promoter in pL117 and the deletion muta-

TABLE 1 Comparative analysis of the transcriptional levels of the most relevant genes (HFX_1558 to HFX_1565) in H. mediterranei DF50 and the�glpR mutant during fructose induction, using a microarray assay

Gene Annotation

DF50 �glpR mutant

Fold change (mean SD) q value (%) Fold change (mean SD) q value (%)

HFX_1558 Fructose-1,6-bisphosphate aldolase 0.82 0.14 4.23 0.86 0.11 10.58HFX_1559 PTS IIB component 29.74 7.31 0 0.92 0.02 12.77HFX_1560 PTS enzyme I 34.97 14.17 0 0.88 0.03 7.85HFX_1561 PTS protein HPr 31.3 12.77 0 0.85 0.17 13.33HFX_1562 PTS IIA component 40.37 16.45 0 0.94 0 20.2HFX_1563 PTS IIC component 31.97 4.33 0 1.11 0.08 17.89HFX_1564 1-Phosphofructokinase 17.33 5.74 0 0.91 0.12 21.9HFX_1565 GlpR family regulator 21.07 2.42 0

FIG 2 GFP expression profiles of the DF50 and �glpR strains harboringsmRSGFP-based reporter plasmids with or without fructose induction. Theplasmids pL117, pPR, and pPF were constructed to express smRSGFP with thepromoters PPTS, PglpR, and Pfba, respectively. Another construct, pL117CR, wastransformed into the �glpR strain to express GlpR (using its native promoter,PglpR) and smRSGFP (using PPTS). The fluorescence intensity was detected asdescribed in the text. Fructose was added to the cell cultures to a final concen-tration of 50 mM (Fru�). TBSL buffer was used as a negative control (Fru�).At least three independent experiments were carried out, and each experimentconsisted of three replicates.

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tions in pL93, pL77, and pL56 exhibited similar basal transcrip-tion activities when cells were grown in AS-168SYL medium (datanot shown). However, in the presence of fructose, the pL56 mu-tant completely lost the ability to respond to the fructose induc-tion. The pL93 and pL77 mutants showed a 2- to 3-fold increase influorescence intensity after the fructose induction. When the pu-tative BRE and TATA box of PPTS were deleted (pL24), the tran-scription activities under both the basal and fructose-inducingconditions were almost undetectable (Fig. 4A). These results re-vealed that the pL77 mutant still contains the main cis-acting ele-ments that respond to fructose induction. Thus, the promoterregion from bp �79 to �16 of pL117 was analyzed using site-directed scanning mutagenesis to pinpoint the essential regionsthat account for the fructose induction. The resulting constructswere named pM7975 to pM�1216, in which the numbers indicatethe mutation region. (For example, pM7975 indicates mutagene-sis from bp �79 to �75 relative to the transcription start site[TSS] of PPTS, and pM�1216 indicates mutagenesis from bp �12to �16. These plasmids were transformed into H. mediterraneiDF50 to generate the reporter strains M7975 to M�1216 for thedetection of fluorescence intensity.

It was observed that mutations in the putative BRE (bp �36 to�31) and TATA box (bp �30 to �25) regions (M3935, M3430,and M2925) and the �10 region (M1410) led to a complete loss oftranscriptional activity. Only three mutants, M7975, M6864, andM2420, had the same response to fructose induction as the pL117transformant, and a 1.5- to 2-fold induction was detected in mu-tants M6360, M5553, M4440, and M1915 (Fig. 4B). It is notewor-thy that the basal fluorescence intensity of M1915 rose to a veryhigh level (19 times that of DF50 harboring pL117). However, thefluorescence intensity of M1915 was still induced by fructose, andthus the region from bp �19 to �15 might not be directly in-volved in fructose activation. GlpR seemed unrelated to any inhi-bition at the region from bp �19 to �15, since the fluorescenceintensity of the �glpR strain harboring pL117 (with or withoutfructose induction) was similar to the basal intensity of DF50 cellsharboring pL117 (without fructose induction) (Fig. 2). These re-sults showed that the knockout of GlpR cannot enhance the PPTS

activity to as high of a level as that detected in the PPTS-mutated

plasmid pM1915. We speculate that either the site from bp �19 to�15 is required for the binding of an unknown inhibitor or themutation from bp �19 to �15 changes the promoter architecture,both of which may lead to a higher activity of the mutated pro-moter under basal conditions.

Notably, the fructose induction did not significantly changethe transcriptional levels of the mutants M7469, M5957, M5755,and M5351 to M4745 (the fold change for each was no more than1.1), indicating that the corresponding regions in these mutantsare important for the fructose-induced upregulation of the PTS.These results revealed that promoter regions I (bp �74 bp to�69), II (bp �59 to �56), and III (bp �52 to �45) are particu-larly important for fructose induction in H. mediterranei, and mu-tations in these three regions made PPTS lose its ability to respondto the fructose induction (Fig. 4B). This phenomenon, which wassimilar to that observed in the �glpR mutant strain harboringpL117 (Fig. 2), indicated that regions I, II, and III were essentialfor the cellular responses to fructose induction and were likely tobe the GlpR binding sites under fructose induction. Interestingly,after analyzing the PPTSs of all six haloarchaea that possess the PTSgene cluster, a conserved 8-bp motif, which overlapped with sevenbase pairs of region III, was identified (Fig. 5). In addition, a pal-indromic DNA sequence pattern belonging to regions I and II,named motif P (short for “palindromic”) in this study, was alsodetected upstream of the 8-bp motif (Fig. 5), implying that theregulatory mechanism of the PTS revealed in H. mediterranei maybe shared by other haloarchaea.

It is noteworthy that the basal fluorescence intensity of mutantM�911 was more than 2.4-fold higher than that of the pL117transformant, and both the basal and induced fluorescence inten-sities in the mutants M�38 and M�1216 decreased to a very lowlevel in these strains (Fig. 4B). These results suggest that the mu-tations in the 5= UTR altered either the mRNA stability or thetranslation efficiency of the PTS gene transcripts.

Generation of a PTS transcript with a shorter 5=UTR due toposttranscriptional processing. The results of scanning mu-tagenesis showed that only one TATA box, BRE, and �10 elementwere identified in the PTS operon (Fig. 4B), which indicated thatthere is only one promoter for PTS transcription. However, twotranscripts with 5=UTRs of different lengths (17 nt and 8 nt) wereobserved among the PTS transcripts (Fig. 1B and C), and the 5=UTR of the PTS gene cluster was found to be important for theirexpression (Fig. 4B). To investigate how the two transcripts con-taining 5= UTRs of different lengths were produced and whetherthey were involved in PTS activation, the features and function ofthis 5= UTR region were further analyzed.

First, RNA folding and the general features of the 17-nt 5=UTRwere predicted using Sfold software, and a stem-loop structurewas indicated. The start site (G) (�10) of the 8-nt 5= UTR waslocated at the loop region (Fig. 6A). DF50 strains harboring dif-ferent plasmids were used to conveniently characterize the 5=UTRat both the transcriptional and translational levels (Fig. 6B). In aprimer extension assay, when the nucleotides in the region frombp �3 to �11 (M�911 and M�38) of the 5=UTR were mutated,the small extension product was hardly detectable (Fig. 6B).Therefore, the production of the shorter GFP gene transcriptslikely depends on the sequence or structure of the 5=UTR (same asthe 5= UTR of PTS transcripts) of the mRNA.

To test this hypothesis, four promoter- or 5= UTR-swappedconstructs were generated, as shown in Fig. 7A. The plasmids,

FIG 3 ChIP-qPCR data suggest that GlpR binds to DNA directly. The relativeenrichment ratio of a 119-bp region of the PPTS (FPTS) immunoprecipitated byGlpR-Myc compared to randomly sheared chromosomal DNA (input sam-ples), using a 145-bp region in 16S rRNA (F16S) as an internal control tonormalize the data, is shown. Enrichments are also compared for a 125-bpcoding region (Fiic) of the ptsC gene and a 189-bp promoter region (FphaE) ofphaE that was not regulated by fructose induction in the microarray data. Cellsamples for the ChIP assay were harvested under growth conditions with (Fru)or without (CK) fructose induction.

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pT8, pT17, and pHSP contained the hsp5 promoter from Halo-bacterium sp. strain NRC-1 (Phsp5) (36) and different 5= UTR re-gions either from PTS (pT8 and pT17) or hsp5 (pHSP), respec-tively, whereas pUTR-M (containing the PTS promoter and the 5=UTR of hsp5) was constructed in a manner similar to that forpHSP by only replacing Phsp5 with PPTS. The results showed that

the two constructs that contained the 17-nt 5=UTR, pT17 and thepositive-control pL117, could both produce two transcripts (cor-responding to T8 and T17 of the PTS transcripts and named T8G

and T17G for the GFP gene transcripts) with different 5=UTRs (Fig.7B), despite the fact that the transcription of these constructs wascontrolled by different promoters (Phsp5 and PPTS, respectively). In

FIG 4 Deletion analysis and site-directed mutagenesis of the PPTS region. (A) Schematic representations (not to scale) of constructs pL117, pL93, pL77, pL56,and pL24. The wild-type PPTS and truncated 5= flanking sequence promoter mutants (solid lines, bp �93 to �17 to �24 to �17) were fused with the smRSGFPreporter gene (gray arrow). (B) Site-directed mutagenesis from bp �79 to �16 of the PPTS. The DNA sequence of wild-type PPTS is shown at the top (pL117). Thetwo transcripts of the reporter gene with 5= UTRs starting from bp �1 and �10, which are the same as observed in the PTS transcripts, were identified byCR-RT-PCR. The mutated nucleotides of different mutants (M7975 to M�1216) are shown below the wild-type (WT) promoter sequence. The basal (nonin-duced) and fructose-induced transcriptional activities of these promoters as revealed by fluorescence intensity were detected using a microplate reader. The basaltranscriptional activities are expressed as a percentage of the pL117 activity (set to 100%), and the fructose-induced fold changes were calculated using thefluorescence intensity. Mutants that cannot respond to fructose are indicated with an asterisk, and mutants with a fluorescence intensity that was hardlydetectable are marked by a dash. The significant fold changes in strains M�38 and M�1216, marked with a rhombus, were caused by the large decreases in thebasal fluorescence intensity. At least three independent experiments were carried out, and each experiment consisted of three replicates.

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contrast, only one transcript was generated from pHSP or pT8, asexpected (Fig. 7B), and the PTS promoter (PPTS) combined withthe hsp5 5=UTR sequence (pUTR-M) was also unable to generatethe shorter transcript. These results indicated that the productionof the transcript with shorter 5= UTR was related only to the se-quence of the longer 5=UTR, and not to the promoter, and there-fore indicated a potential posttranscriptional processing of PTStranscripts.

Physiological significance of the generation of the shorter 5=UTR. To understand the physiological significance of the produc-tion of two transcripts containing 5=UTRs of different lengths, thetranslation efficiencies of these two transcripts were investigated.The relative amounts of GFP gene transcripts (detected by qRT-PCR) and their translation activities (measured by fluorescenceintensity) were determined using DF50 strains harboring pT8 orpT17 (Fig. 8A). There was no significant difference in the amountof GFP gene transcripts between DF50 strains harboring pT8 orpT17 as evaluated by the Student t test, but the GFP expressionfrom the shorter transcript in the DF50 strain harboring pT8 wasover 6-fold higher than the GFP expression from the mixture oftwo transcripts in the DF50 strain harboring pT17 (Fig. 8A). Theseresults indicated that the translation efficiency of the shorter tran-script was much higher than that of the longer transcript.

Further research was performed to find a correlation betweenthe fructose induction and the production of the two transcripts.The primer extension products of the DF50 and �glpR strainsharboring pL117 with or without fructose induction were ana-lyzed. The results showed that the amounts of the two transcriptsincreased as a result of fructose induction in the DF50 strain, andthe ratio of the shorter transcript to the longer transcript was alsoincreased (Fig. 8B). To confirm this result quantitatively, CR-RT-PCR was performed on the DF50 and �glpR strains with or with-out fructose induction. The ratio of clone counts of the PTS tran-script with the 8-nt 5=UTR to that of the transcript with the 17-nt5= UTR was doubled (from 14% to 31%) when DF50 cells wereinduced by fructose (Table 2). The ratios of T8 to T17 counts in the�glpR strain with or without induction were similar to each otherand much closer to the noninduced ratio in the DF50 strain (Table2). These results indicate that the translation efficiency of the PTSgenes would be enhanced when the cells were induced by fructosedue to the increased proportion of the shorter transcript.

F-1-P may acts as a positive intracellular effector. During PTSregulation, the derivatives of sugars always serve as positive ornegative effectors to enhance or repress the activity of regulators.F-1-P was shown to act as an important intracellular effector forthe transcriptional regulation of the PTS in many bacteria (2, 14,

FIG 5 Multiple alignments of promoter sequences of the PPTSs in the haloarchaea which contain at least one complete PTS. Bases marked with asterisks are thesequences of regions I, II, and III. A palindromic DNA sequence (underlined, motif P) and an 8-bp sequence (boxed) are indicated. These sequences were foundto be conserved via the alignment of the PPTS in all promoter regions of haloarchaeal PTS.

FIG 6 Mutagenesis analysis of the 5=UTRs of PTS transcripts. (A) In silico-predicted secondary structure of the 17-nt 5=UTR of mRNA, (�G°37 � �2.50). Thedesigned mutation regions are marked with lines. (B) Electrophoretic analysis of the primer extension products of the GFP gene in strains M�911, M�38, andM1�2 after induction by fructose. DF50 cells harboring pGFP-0 were used as a negative control (NC), and DF50 cells harboring pL117 were used as a positivecontrol (PC). The 59-nt and 41-nt oligonucleotides were used as molecular markers. The nonspecific products appearing in the NC are indicated with a and b,and the two primer extension products, corresponding to the transcript with the 17-nt (T17G) or 8-nt (T8G) 5= UTR, are indicated with arrows.

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15, 37). To investigate the function of F-1-P during PTS regula-tion, the smRSGFP fusion reporter assay was performed with thepL117 transformant of H. mediterranei. In the fruK knockout mu-tant containing pL117, whether induced by fructose or not, thefluorescence intensity increased to a very high level (more than 20times higher than that in DF50) and the induced expression ofPTS by fructose disappeared (Table 3). In H. volcanii, it has beenrevealed that fructose is transported through the PTS, whichwould generate F-1-P, after which it is further phosphorylated by

1-PFK (encoded by fruK) (12). The high level of GFP gene expres-sion in H. mediterranei �fruK may be caused by the accumulationof F-1-P when 1-PFK is inactivated, which implies that F-1-P mayenhance the expression of the PTS as an intracellular effector asobserved in bacteria. This hypothesis is also supported by the flu-orescence intensity of H. mediterranei DF50 cells harboring pL117when F-1-P is added to the culture medium (see Fig. S3 in thesupplemental material). Either fructose or F-1-P induction signif-icantly increased the fluorescence intensity, whereas the other de-rivative of fructose, fructose-1,6-bisphosphate (F-1,6-2P), de-creased the fluorescence intensity (see Fig. S3 in the supplementalmaterial).

DISCUSSION

During fructose induction, GlpR has been shown to be an indis-pensable activator for the upregulation of the fructose-specificPTS gene cluster (Table 1; Fig. 2) through direct binding to PPTS inH. mediterranei (Fig. 3). Therefore, GlpR is essential for the cellu-lar responses to fructose induction. Interestingly, in H. volcanii,GlpR has been shown to be a global regulator by repressing thetranscription of the key enzymes, including KDGK and PFK,when using glycerol as the carbon source (13). It seemed that thefunction of GlpR was decided by the environmental carbonsources. Through the regulator GlpR, glycerol represses sugar me-tabolism and fructose activates PTS expression. Previous reportssuggest that the DeoR-type proteins always contain several highlyconserved regions, one of which is the second helix of the helix-turn-helix (HTH) DNA binding motif in the N terminus (38). Theother conserved regions are involved in oligomerization or in-ducer binding (in many cases, the inducer is a phosphorylatedsugar). As one of the DeoR-type proteins, it makes sense that GlpRcould be activated through the interaction of its C-terminal sensor

FIG 7 Primer extension assay for identifying the different transcripts generated by Phsp5-directed or PPTS-directed reporter genes. (A) Sequences of constructspT8, pT17, pHSP, and pUTR-M. The TSS (G) of Phsp5 or the TSS (A) of PPTS in the four constructs is marked with asterisks, and the different-length 5=UTR ofeach construct is underlined. The sequence of the extension primer is boxed, and the expected size of the extension product is indicated in parentheses. Thedifference between pT8, pT17, and pHSP is the downstream sequence of the TSS (G), which is the 8-nt (pT8) or 17-nt (pT17) 5=UTR sequence of the PTS or the5= UTR sequence of the wild-type hsp5 (pHSP), respectively. The only difference between pHSP and pUTR-M is that the Phsp5 promoter is replaced by PPTS inpUTR-M. (B) The primer extension products of pT8, pT17, pHSP (T56), and pUTR-M were analyzed by electrophoresis. The primer extension products withsame size are indicated with arrows (T56, T17G, or T8G).

FIG 8 Functional characterization of the in vivo generation of the shorter 5=UTR in H. mediterranei using a reporter gene. (A) The relative level of tran-scription activity and the translation efficiency of GFP gene transcripts in DF50strains harboring pT8 or pT17 were determined via qRT-PCR and a fluores-cence reporter assay. The levels of GFP gene transcription and the fluorescenceintensity in strain DF50 harboring pT17 were both assigned a value of 1. Atleast three independent experiments were carried out, and each experimentconsisted of three replicates. The statistical significance of the difference be-tween the DF50 strains harboring pT8 or pT17 was analyzed using the Studentt test. (B) Electrophoretic analysis of the primer extension products of the GFPgene in DF50(pL117) and �glpR(pL117) strains with (�) or without (�)fructose induction. Major transcripts are indicated on the left as described forFig. 6.

TABLE 2 Statistical results for the shorter and longer PTS transcripts instrain DF50 and the �glpR mutant with or without fructose induction

StrainFructoseinduction

Counts

Ratio (8-nt transcript/17-nt transcript), %

8-nttranscript

17-nttranscript

DF50 � 5 36 14� 8 26 31

�glpR mutant � 2 14 14� 2 20 10

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domain with the fructose effector F-1-P in H. mediterranei. Theactivated GlpR (or GlpR accompanied by unknown regulators)could then bind to the promoter region through the N-terminalDNA binding motif to increase the transcriptional activity of PPTS.This mode of action of GlpR was supported by the study of an-other DeoR-type transcriptional regulator, SugR, in Corynebacte-rium glutamicum (15, 39–41). Although the molecular details ofthe activation by GlpR require further investigation, one hypoth-esis, based on other transcription activation models in archaea(42–44), is that the activated GlpR recruits general transcriptionfactors (transcription factor Bs [TFBs] and TATA binding pro-teins [TBPs]) to bind to the TATA box or BRE to enhance tran-scription.

According to a previous report, the possible binding sites of theGlpR as a repressor are located at the BRE or downstream of theTATA box as determined by in silico searching for the invertedrepeat sequence (13). However, this kind of binding site that wasreported for H. volcanii GlpR was not observed in the PTS pro-moter region in H. mediterranei. Instead, an upstream sequence(including regions I, II, and III) of PPTS was indicated as the pos-sible binding site of GlpR by the mutation scanning experiment(Fig. 4B). This finding showed a positional similarity with theupstream activator sequence (UAS) (from bp �52 to �39) of thebop gene in Halobacterium sp. strain NRC-1 (44–46). The con-served motifs (motif P and the 8-bp motif) in promoter regions I,II, and III of the PTS implied that the mechanism of PTS regula-tion by GlpR is similar in haloarchaea (Fig. 5).

It is noteworthy that the shorter PTS transcript would be gen-erated through posttranscriptional processing in H. mediterranei(Fig. 7). This type of expression seemed to be different from itsbacterial counterpart. Multiple TSSs were reported in the PTSgenes of E. coli, which resulted from multiple promoters upstreamof the coding sequences of the PTS genes and were influenced byDNA supercoiling and the transcription factor cAMP receptorprotein (47), but the translation efficiencies of different transcriptpatterns were not very clear. Further analysis of the 5= UTR se-quences of T8G and T17G indicated that the relatively more efficienttranslation of T8G was probably due to the presence of the shorter5= UTR (Fig. 8A). This hypothesis that the length of the 5= UTRaffects the translation efficiency of mRNA has also been givenfor other haloarchaea. In Halobacterium salinarum, leaderlessmRNAs showed a higher translation activity than mRNAs with theShine-Dalgarno (SD) sequence (48). On the other hand, the pre-dicted stem-loop structure in T17 (Fig. 6A) might also inhibit thereorganization or binding of the ribosome and hence repress thetranslation. It can be speculated that when induced by fructose,the translation efficiency of the PTS mRNA could be enhanced by

increasing the ratio of T8 to T17 (Table 2). The relatively higherproportion of T17 under conditions without fructose implied aconstitutively low-level expression of the PTS.

The fructose-specific PTS of haloarchaea was thought to beacquired from bacteria by horizontal gene transfer (HGT) duringevolution (12, 49). As a “gift from the neighbors,” the haloarchaealPTS was also capable of “borrowing” the regulatory mechanismfrom bacteria at the transcriptional level. Furthermore, to accli-mate to nutrient fluctuations in a competitive extreme hypersalineenvironment, the haloarchaea evolved their own mechanisms tocontrol the PTS at the posttranscriptional and translational levels.Based on the results in this study and previous reports, we proposea working model for PTS regulation in haloarchaea (Fig. 9). In thismodel, fructose is transported into the cell and phosphorylated toF-1-P via the PTS and is further catalyzed to F-1,6-2P by 1-PFK.GlpR (or a combination of GlpR and other, unknown regulators)upregulates the transcription of this PTS gene cluster after theinduction by fructose via direct binding to the PPTS, most probablyat the three conserved regions. F-1-P may act as the intracellularinducer, while F-1,6-2P may act as the negative effector, to beinvolved in this transcriptional regulation of PTS gene expression.A posttranscriptional processing of the PTS transcripts at the 5=UTR, which increases the translational efficiency, is also involvedin the PTS activation upon fructose induction (Fig. 9).

In conclusion, the activation at the both transcriptional andtranslational levels would make the haloarchaeal PTS more effi-cient in response to environmental fructose. Although the work-ing model has explained the main mechanisms of PTS regulationin haloarchaea, further study is warranted to determine whetherother transcriptional regulators are involved in the regulation ofPTS expression and to elucidate the mechanism of posttranscrip-tional processing. Such studies would help toward a comprehen-sive understanding of PTS regulation in haloarchaea.

ACKNOWLEDGMENTS

We thank Julie A. Maupin-Furlow (University of Florida, USA) for pro-viding us with plasmid pJAM1020.

This work was supported by grants from the National Natural ScienceFoundation of China (31330001, 30925001, and 31271334) and the Chi-nese Academy of Sciences (KSCX2-EW-G-2-4).

TABLE 3 Expression of the smRSGFP fusion reporter gene in H.mediterranei strains with or without fructose inductiona

Exptno.

Relevant hostgenotype

Relevantplasmid

Fluorescence intensity(mean SD)

FoldchangeNoninduced

Fructoseinduced

1 DF50 pL117 896 158 5,052 753 5.642 �fruK pL117 22,108 570 19,652 612 0.89a At least three independent experiments were carried out, and each experimentconsisted of three replicates.

FIG 9 A working model of the regulation of PTS expression and fructoseutilization in H. mediterranei. CM, cytoplasmic membrane; 1-PFK, 1-phos-phofructokinase; F-1-P, fructose-1-phosphate; F-1,6-2P, fructose-1,6-bispho-sphate; TIC, transcriptional initiation complex.

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