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Specific features of l-histidine production by Escherichia coli concerned with feedback control of AICAR formation and inorganic phosphate/metal transport

  • Evgeniya A. Malykh1,
  • Ivan A. Butov1,
  • Anna B. Ravcheeva1,
  • Alexander A. Krylov1,
  • Sergey V. Mashko1 and
  • Nataliya V. Stoynova1Email author
Microbial Cell Factories201817:42

Received: 12 December 2017

Accepted: 8 March 2018

Published: 15 March 2018



In the l-histidine (His) biosynthetic pathway of Escherichia coli, the first key enzyme, ATP-phosphoribosyltransferase (ATP-PRT, HisG), is subject to different types of inhibition. Eliminating the feedback inhibition of HisG by the His end product is an important step that enables the oversynthesis of His in breeding strains. However, the previously reported feedback inhibition-resistant mutant enzyme from E. coli, HisGE271K, is inhibited by purine nucleotides, particularly ADP and AMP, via competitive inhibition with its ATP substrate. 5-Aminoimidazole-4-carboxamide ribonucleotide (AICAR), which is formed not only during His biosynthesis but also during de novo purine biosynthesis, acts as a natural analog of AMP and substitutes for it in some enzymatic reactions. We hypothesized that AICAR could control its own formation, particularly through the His biosynthetic pathway, by negatively influencing HisG enzymatic activity, which would make preventing ATP-PRT transferase inhibition by AICAR crucial for His overproduction.


For the first time, both the native E. coli HisG and the previously described feedback-resistant mutant HisGE271K enzymes were shown to be sensitive to inhibition by AICAR, a structural analog of AMP. To circumvent the negative effect that AICAR has on His synthesis, we constructed the new His-producing strain EA83 and demonstrated its improved histidine production. This increased production was particularly associated with the improved conversion of AICAR to ATP due to purH and purA gene overexpression; additionally, the PitA-dependent phosphate/metal (Me2+-Pi) transport system was modified by a pitA gene deletion. This His-producing strain unexpectedly exhibited decreased alkaline phosphatase activity at low Pi concentrations. AICAR was consequently hypothesized inhibit the two-component PhoBR system, which controls Pho regulon gene expression.


Inhibition of a key enzyme in the His biosynthetic pathway, HisG, by AICAR, which is formed in this pathway, generates a serious bottleneck during His production. The constructed His-producing strain demonstrated the enhanced expression of genes that encode enzymes involved in the metabolism of AICAR to ATP, which is a substrate of HisG, and thus led to improved His accumulation.


l -HistidineInorganic phosphate/metal transportATP-phosphoribosyltransferaseAICAR (ZMP)PitAPho regulon Escherichia coli l-Histidine production


l-Histidine (His) is an essential amino acid that is used in the pharmaceutical industry as a component of nutritious mixes for infant and adult humans [1]. In biochemical studies, His is a growth factor involved with many primary metabolites [2, 3]. Industrial His-producing strains of Corynebacterium glutamicum, Brevibacterium flavum, Serratia marcescens and Escherichia coli have been described [3].

The biosynthesis of His was initially well characterized in Salmonella typhimurium and E. coli as imperative model systems to study the fundamental processes connected with this function, including transcriptional attenuation, gene expression, and enzymatic feedback regulation [46].

In bacterial cells, the His biosynthesis pathway is associated with ten biochemical reactions and nine enzymes that contribute to the conversion of two biosynthetic precursors, phosphoribosyl pyrophosphate (PRPP) and adenosine triphosphate (ATP), into this amino acid (Fig. 1) [6].
Fig. 1

Metabolism of AICAR to ATP is an essential step in His synthesis. Prs ribose-phosphate diphosphokinase, ATP adenosine triphosphate, R5F ribose 5-phosphate, PRPP phosphoribosyl pyrophosphate, PR-ATP phosphoribosyl-ATP, PR-AMP phosphoribosyl-AMP, PR-F-AICAR-P phosphoribosylformimino-AICAR-phosphate, PR-FAR phosphoribulosylformimino-AICAR-phosphate, AICAR 5-aminoimidazole-4-carboxamide ribonucleotide, FAICAR 5′-phosphoribosyl-5-formamido-4-imidazole carboxamide, IMP inosine 5′-phosphate, AS adenylosuccinate, AMP adenosine monophosphate, ADP adenosine diphosphate, IGP imidazole glycerol phosphate, IA-P imidazole acetol-phosphate, His-ol-P l-histidinol-phosphate, His-ol histidinol, Histidine l-histidine, HisG ATP-phosphoribosyltransferase, HisIE bifunctional enzyme (HisE phosphoribosyl-ATP pyrophosphatase, HisI phosphoribosyl-AMP cyclohydrolase), HisA phosphoribosylformimino-5-amino-1-phosphoribosyl-4-imidazole carboxamide isomerase, HisHF IGP synthase, HisB bifunctional imidazoleglycerol-phosphate dehydratase/histidinol-phosphatase, HisC histidinol-phosphate aminotransferase, HisD bifunctional histidinal dehydrogenase/histidinol dehydrogenase, PurH bifunctional AICAR transformylase/IMP cyclohydrolase, PurA adenylosuccinate synthetase, PurB adenylosuccinate lyase, Adk adenylate kinase, Gln glutamine, Glu glutamate, fTHF formyltetrahydrofolate, THF tetrahydrofolate, Asp aspartate, GTP guanosine-triphosphate, GDP guanosine-diphosphate, PP i inorganic pyrophosphate, P i inorganic phosphate, NAD + /NADH nicotinamide adenine dinucleotide oxidized/reduced form, fum fumaric acid. Feedback inhibition by His is indicated by red line; competitive inhibitions by ATP-PRT, AMP and ADP are indicated by dot lines; suggested competitive inhibition by AICAR is indicated by the red dot line

The first key step in the His biosynthetic pathway, the condensation of PRPP and ATP to form phosphoribosyl-ATP (PR-ATP), is catalyzed by the ATP-PR transferase (HisG, EC, which is exposed to a variety of types of inhibition: allosteric feedback inhibition by His, competitive inhibition by adenosine mono- and diphosphates (AMP and ADP, respectively) [7], and competitive product inhibition by PR-ATP. Eliminating the His-mediated feedback inhibition of HisG plays a critical role in the regulation of His biosynthesis [6, 8]. Feedback-resistant HisG-mutant enzymes from S. typhimurium [9] and C. glutamicum [10, 11] have been previously characterized. The HisGE271K, Fbr-mutant from E. coli was obtained by traditional selection methods several decades ago [12], and its properties were later investigated [13]. Despite having complete resistance to feedback inhibition by His, this mutant enzyme is still susceptible to competitive inhibition by purine nucleotides, particularly ADP and AMP [12]. This finding suggests that HisGE271K, similar to the native HisG, could be the target of inhibition by the AMP structural analog, 5-aminoimidazole-4-carboxamide-1-beta-d-ribofuranosyl 5′-monophosphate (AICAR). Figure 1 indicates that AICAR is formed via the His biosynthetic pathway as well as through the de novo purine biosynthetic pathway [6, 14]. In the present study, we confirmed that both the native (HisGWT) and feedback-resistant (HisGE271K) enzymes were sensitive to inhibition by AICAR. The formation of AICAR through the His biosynthetic pathway in E. coli is thus negatively controlled by end-product (AICAR) inhibition of the first key enzyme in the pathway, HisG. Considering that His biosynthesis is accompanied by equimolar AICAR generation, the effective metabolism of AICAR to ATP, which in turn functions as the HisG substrate, appears crucial for the production of this amino acid.

The reactions that convert AICAR into ATP include the use of bi-functional PurH (EC + EC [15], PurA (EC [16], PurB (EC, Adk (EC and H+-ATP synthase (F1F0-ATP synthase) [17] (Fig. 1). Since the final step of ATP synthesis from AICAR includes ADP phosphorylation by inorganic phosphate (Pi) that is catalyzed by the proton force-dependent ATP synthase, the maintenance of intracellular Pi at the appropriate level is a significant requirement for the efficiency of the whole pathway.

The two major Pi transport systems were initially described in E. coli cells: Pst (PstSCAB, phosphate-specific transport) and Pit (PitA, phosphate inorganic transport) [1821]. PstSCAB, a member of the ABC superfamily of ATP-dependent transporters, is the major high-affinity, low-velocity phosphate uptake system. Under conditions of Pi limitation, the Pst system is activated more than 100-fold, and Pi is primarily taken up by the Pst transporter. In addition, at basal expression levels, the Pst system and not PitA (see below) primarily contributes to Pi uptake when excess Pi is present [22]. E. coli also possesses a high-velocity, low-affinity Pi transporter, PitA, which does not belong to the phosphate (Pho) regulon; PitA is dependent on the proton motive force for energization, and P i uptake via PitA is defined by the presence of divalent cations such as Zn2+, Mg2+, Ca2+, Co2+, and Mn2+, which form soluble metal phosphate (MeHPO4) complexes [23]. PitA gene expression was reported to be modulated in a chemically defined medium by the availability of Pi and Zn2+ ions [24], and PitA functions primarily as a transporter of divalent metal cations (Zn2+) complexed with Pi [22, 25]. A third Pi transport gene, pitB, encodes a functional Pi transporter that is a homolog of PitA, sharing 81% identity (, BLAST® data); pitB is repressed at low Pi levels by the Pho regulon [26] and likely does not play a role in Pi uptake in normal cells because it is not expressed under normal growth conditions [22]. Two additional transporters, encoded by glpT and uphT, transport Pi with low affinity [2729]; however, in the absence of Pst and Pit, these systems cannot support growth when phosphate is provided as Pi [24, 30]. Maintaining a sufficient Pi pool is extremely important for many energy-consuming cellular processes [18], particularly for His synthesis.

Oversynthesis of any cell metabolite, e.g., His, alters the carbon fluxes and pools of intermediate compounds, possibly resulting in consequences that are generally unpredictable. In the present study, some effects were revealed to be related to His overproduction by E. coli, and the elucidation of these effects has begun. AICAR negatively influenced His biosynthesis in E. coli, and a strategy for the efficient conversion of this intermediate to ATP by enhancing the de novo purine biosynthetic pathway genes, purH and purA, was developed that improved His production. The influence of AICAR on the activity of HisG was studied and revealed that AICAR is a structural analog of AMP; AICAR was even a stronger inhibitor of HisG (Ki = 0.65 mM) than AMP (Ki = 2.15 mM). His oversynthesis was accompanied with a change in the functioning of the Pho regulon that was explained by changes in the AICAR pool. Furthermore, pitA gene deletion positively influenced His overproduction, and although the precise mechanism underlying this effect remains unclear, several explanations based on biological role of PitA as a Me2+-Pi importer were supposed.


Strains, plasmids and media

All bacterial strains and plasmids used in this study are listed in Table 1.
Table 1

E. coli strains and plasmids used in this study

Strain or plasmid





Escherichia coli K-12 wild-type

VKPMa B-6195


E. coli B F ompT gal dcm lon hsdS B (r B m B ) λ(DE3 [lacI PlacUV5-T7gene1 ind1 sam7 nin5]) [malB+]K–12S)



MG1655+-[ΔpurR P his hisLhisGE271KDCBHAFI]


 MG1655 [ΔpurR::CmR]

MG1655 wild-type E. coli K-12, but ΔpurR::λattR-catattL

Laboratory collection

 BW25113 [ΔpurH::KmR]

lacIq rrnBT14 ΔlacZWJ16 hsdR514 ΔaraBADAH33 ΔrhaBADLD78, ΔpurH::FRT-kan-FRT


 MG1655 [Δ(φ80-attB)]

MG1655 with deleted native (φ80-attB) site


 MG1655 [Δ(φ80-attB) IS 5.11::φ80-attB]

MG1655 with deleted native φ80 attB site and reconstruction of attB site in IS 5.11 locus


 MG1655 [Δ(φ80-attB) IS 5.11::CmR-Ptac21-purA]

MG1655-[Δ(φ80-attB) IS 5.11::φ80-attB], but IS 5.11::(λ-attB) Ptac21-purA

This study

 MG1655 [CmR-PL-purH]

MG1655, but λattR-catattL-PL-purH

This study

 MG1655 [ΔpitA::KmR]

MG1655, but ΔpitA::λattR-kanattL

This study

 MG1655 [CmR-PLtac-pitA]

MG1655, but λattR-catattL-PLtac-pitA

This study

 MG1655 [ΔpurR::CmR ΔpurH:: KmR]

MG1655, but ΔpurR::λattR-catattL and ΔpurR::FRT-kan-FRT

This study

 KF37 [ΔpitA::KmR]

KF37, but ΔpitA::λattR-kanattL

This study

 KF37 [IS5.11::CmR-Ptac21-purA pitA

KF37, but IS 5.11:: λattR-catattL Ptac21-purA pitA

This study

 KF37 [IS5.11::CmR-Ptac21-purA pitA+]

KF37, but IS 5.11::λattR-catattL Ptac21-purA pitA+

This study


KF37, but IS 5.11::(λ-attB) Ptac21-purA pitA

This study


EA79, but (λ-attB) PL-purH

This study

 CC118 λpir+

Host strain for maintenance of pir-dependent recombinant plasmids


 MG1655 [Δ(φ80-attB)  ∆yibH::φ80-attB]

MG1655 with deleted native φ80 attB site and reconstruction of attB site instead of yibH locus

This study

 MG1655 [∆yibH::TcR-phoBDBD]

MG1655 [Δ(φ80-attB) ∆yibH::φ80-attB], but ∆yibH::λattR- tetARattL-phoBDBD

This study

 MG1655 [∆yibH::KmR-PlacUV5-phoBDBD]

MG1655 [Δ(φ80-attB) ∆yibH:: λattR-tetARattL -phoBDBD], but ∆yibH::λattR-kanattL-PlacUV5-phoBDBD

This study

 KF37 [∆yibH::TcR-phoBDBD]

KF37, but ∆yibH::λattR- tetARattL-phoBDBD

This study

 KF37 [∆yibH::KmR-PlacUV5-phoBDBD]

KF37, but ∆yibH::λattR-kanattL-PlacUV5-phoBDBD

This study



oriR101, repA101ts, araC, P araB -[γ, β, exo of phage λ], ApR; used as a donor of λRed-genes to provide λRed-dependent recombination



oriR101, repA101ts, λcIts857, λPR→λxis-int, ApR; used as a helper plasmid for thermoinducible expression of the λ xis-int genes



oriR101, repA101ts, λcIts857, λPR→φ80-int, ApR; used as a helper plasmid for thermoinducible expression of the φ80-int gene

[37]; GenBank accession number AY048726


ApR, pBR322 origin, P lacI -lacI, and T7 promoter/O lac , T7 transcription start, His6-Tag coding sequence, T7 terminator



oriR101, repA, MCS, ApR, λattR-kanattL—donor of λXis/Int-excisable KmR marker



ApR, pET15b containing hisG gene

This study


ApR, pET15b containing the mutant hisGE271K gene

This study


attP phi80, pAH162, λattL-tetA-tetRattR

[34]; Gene Bank accession number AY048738

 pMW119-P lac -lacI

ApR, low-copy-number vector pMW119, containing P lac -lacI

Laboratory collection

 pMW119-P lac -lacI-purA

ApR, low-copy-number vector pMW119, containing P lac -lacI-purA

This study


oriRγ, φ80-attP, λattL-tetA-tetRattR, purA

This study


PlacUV5-lacI, rep(pMB1), ApR, CmR



oriRγ, φ80-attP, λattL-tetA-tetRattR, promoter-less phoBDBD

This study

Ap R ampicillin resistance, Cm R chloramphenicol resistance, Tc R tetracycline resistance

aVKPM Russian National Collection of Industrial Microorganisms

The following media were used to culture bacteria: Luria–Bertani (LB), M9, SOB, SOC [31], and MOPS, and the MOPS medium was supplemented with 0.25 mM KH2PO4. Glucose (0.4%) was added to minimal media as a carbon source. The antibiotics ampicillin (Ap, 100 mg/L), chloramphenicol (Cm, 20 mg/L), tetracycline (Tc, 20 mg/L), and kanamycin (Km, 50 mg/L) were used when necessary.

Test tube cultivation conditions

For His accumulation, the strains were grown in LB medium at 30 °C overnight; 0.1 mL of each culture was then inoculated into 2 mL of fermentation medium in a test tube, which was then cultivated for 72 h at 30 °C with shaking on a rotary shaker (250 rpm), until all of the glucose was consumed. The composition of the fermentation medium (g/L) was as follows: soybean meal hydrolysate, 0.1 g; l-aspartate, 0.5 g; (NH4)2SO4, 9 g; KCl, 0.5 g; KH2PO4, 0.25 g; MgSO4·7H2O, 0.2 g; FeSO4·7H2O, 0.01 g; MnSO4·5H2O, 0.01 g; ZnSO4·7H2O, 0.01 g; adenosine, 0.1 g; vitamin B1, 0.0005 g; betaine, 1 g; CaCO3, 30 g; and glucose, 25 g, as the carbon source; the pH was adjusted to 6.0.

DNA manipulation

Genetic manipulation of E. coli and techniques for the isolation and manipulation of nucleic acids were performed according to standard protocols [31]. Restriction enzymes, T4 DNA ligase, High Fidelity PCR Enzyme Mix, Taq polymerase, and 1-kb DNA Ladder were purchased from Thermo Scientific Inc. (USA). Plasmids and genomic DNA were isolated using QIAGEN Plasmid Mini Kits (QIAGEN GmbH, Germany) and Bacterial Genomic DNA Kits (Sigma), respectively. QIAquick Gel Extraction kits (QIAGEN GmbH, Germany) were used to isolate DNA from agarose gels. Oligonucleotides were purchased from Evrogen (Russia). The sequences of the oligonucleotide primers are presented in Additional file 1: Table S1.

Quantitative determination of the l-histidine concentration

The amount of His that accumulated in the medium was determined by thin-layer chromatography (TLC) using plates coated with silica gel (Merck, Germany). Samples were applied to the TLC plates using Linomat 5 (Camag, Switzerland). The plates were developed with a mobile phase consisting of propan-2-ol:acetone:25% aqueous ammonia:water = 12.5:12.5:3:2 (v/v). A solution of ninhydrin (1%) in acetone was used as the visualizing reagent; the plates were dried and then scanned at 520 nm using a Linomat 5 scanner (Camag, Switzerland).

pET15b-hisG plasmid construction

The native hisG gene was cloned into a pET15b plasmid vector after PCR amplification (see Additional file 1: Table S1 for details, primers 4, 5). The obtained PCR fragments were treated with BamHI and NdeI endonucleases and ligated into a pET15b vector that had been treated with the same enzymes. The obtained pET15b-hisG plasmid carries the gene encoding HisG with an N-terminal cleavable His6-Tag (HT-HisG) for affinity purification. The obtained plasmid was introduced into the strain E. coli BL21(DE3) for HT-HisG induced biosynthesis followed by tagged-protein purification. Plasmid pET15b-hisGE271K, which harbors the mutant feedback-resistant His6-tagged HisG (HT-HisGE271K) gene, was obtained in a similar manner.

HisG expression and purification

Escherichia coli BL21(DE3)/pET15-hisG was grown in 50 mL of LB medium until the OD560 was 0.8. Protein expression was induced by the addition of isopropyl β-d-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM, which was followed by incubation for 4 h. The cells were centrifuged, washed twice with 50 mL of 100 mM NaCl solution, centrifuged again, resuspended in 50 mL of buffer I (300 mM Tris–HCl, 300 mM KCl, and 1 mM PMSF, pH 8.1) and disrupted by two passages through a cold French press cell at 2000 psi. Unbroken cells and cell debris were removed by centrifugation at 12,000 rpm for 30 min at 4 °C. The supernatant was applied to a 1-mL HiTrap® column (Pharmacia, Sweden); this column was then washed with 10 ml of buffer I, and bound protein was eluted with a linear gradient of buffer I and buffer II (20 mM Tris–HCl and 400 mM imidazole, pH 8). The resulting HT-HisG preparation was further purified by gel filtration using a 10-mL BioGel P10 column (Pharmacia, Sweden) equilibrated with buffer III (20 mM potassium phosphate buffer, pH 7, 1 mM DTT, 10 μM PLP, and 10% (w/v) glycerol). This method was used to purify two ATP-phosphoribosyltransferases (ATP-PRTs): native HT-HisG and mutant HT-HisGE271K. The purity of the obtained proteins was greater than 90% according to SDS-PAGE analysis (see Additional file 2: Figure S1).

ATP-phosphoribosyltransferase assay

The HisG enzymatic activity and initial velocity of the forward phosphoribosyltransferase reaction were measured by monitoring the formation of PR-ATP (ε290 = 3600/M/cm [40]) in the presence of E. coli inorganic pyrophosphatase (PPase, Sigma-Aldrich, USA). Reactions were performed in UV-star 96-well microplates (Greiner Bio-One, Germany). The reaction mixture consisted of 100 mM Tris–HCl (pH 8.1), 100 mM KCl, 10 mM MgCl2, 5 mM ATP, 1 mM PRPP, 10 mU of pyrophosphatase and 500 nM E. coli HisG [41]. The reaction was initiated by the addition of ATP. The absorption at 290 nm was monitored for 30 min in 2-min intervals (Tecan, Switzerland). A reaction mixture containing water instead of ATP substrate was used as a blank control. To test the inhibition of HisG by AMP and AICAR, these compounds were added to the initial reaction mix. Notably, the molar extinction coefficients of AICAR and AMP are ε290, AICAR = 2700/M/cm and ε290, AMP = 240/M/cm. AICAR concentrations higher than 1 mM were therefore undesirable for the measurements, but AMP concentrations as high as 5 mM could be used without significant interference. To obtain the values of KI, AMP and KI, AICAR, AMP or AICAR was added to the reaction mix to a final concentration of 20 and 1 mM, respectively. To calculate the inhibition constants, the following formula was used [42]: K I  = [I]/((K Mi /K M ) − 1), where K I is the inhibition constant, [I] is the concentration of the inhibitor, K M is the Michaelis constant, and K Mi is the K M value in the presence of inhibitor.

Alkaline phosphatase (PhoA) assay

The enzymatic activity of E. coli alkaline phosphatase (the phoA gene protein product, E.C. was measured according to the method of Brickman and Beckwith [43] with modifications. Cells were grown in a flask for 24 h in MOPS medium supplemented with 0.4% glucose and 0.25 mM KH2PO4 and were then washed with 0.9% NaCl. The cells were then lysed by sonication, and cell debris was removed by centrifugation at 12,000 rpm for 20 min at 4 °C. The protein concentration in the supernatant was determined using a standard Bradford Protein assay [44]. PhoA enzymatic activity was analyzed 2 or 24 h after Pi exhaustion. Enzyme reactions were performed in 96-well UV-star microplates (Greiner Bio-On, Germany) and consisted of 500 mM Tris–HCl, pH 8.0, 1 mM MgCl2, and supernatant (or supernatant diluted in 0.9% NaCl) containing from 0.02 to 0.10 mg of protein. p-Nitrophenyl-phosphate (pNPP) was used as the substrate. After pNPP addition, the reaction mixture was incubated at 37 °C for 3–4 min, by which time it had turned yellow. The reaction was stopped by the addition of 1 M KH2PO4, and the absorbance at 410 nm was measured in a UV-star microplate cell against a control reaction that did not contain protein.

Inorganic Pi measurement

The amount of Pi in culture media was determined using a common method based on phosphomolybdate reduction to molybdenum blue [45, 46]. In 96-well microplates containing 0.1 mL of each sample to be analyzed, 0.075 mL of ammonium molybdate-ferrous sulfate (a colored mixture) was added, and the resulting solution was vortexed and incubated at room temperature for 10 min. The colored mixture was prepared daily by mixing 4 vol of 2.5% ammonium molybdate solution with 1 vol of 2.5% ferrous sulfate solution. For the calibration curve, the colored mixture was added to samples containing standard solutions of KH2PO4. The absorbance was measured at 700 nm. A sample without added Pi served as a blank.

Construction of strains and plasmids

Insertions and deletions in the chromosome of E. coli, usually the MG1655K-12 strain, were prepared via λ-Red modification according to the method of Datsenko and Wanner [36], exploiting the λXis/Int system for marker excision [34]. The plasmid pKD46, carrying the arabinose-inducible λ-Red genes [36] and kindly gifted by Dr. Wanner, was used. \(\upvarphi\)80-mediated site-specific integration was carried out according to the method of Haldimann and Wanner with the thermoinducible \(\upvarphi\)80-Int gene in the pAH123 plasmid [37]. Specially designed cassettes with CmR, KmR and TcR markers bracketed by hybrid λattPattB-sites (λattL and λattR) were transferred into E. coli strains by P1-mediated general transduction (P1-duction) [47]. Finally, CmR, KmR and TcR were eliminated from the E. coli chromosome using a λXis/Int site-specific recombination system with pMWts-λInt/Xis as a helper plasmid [34].

Construction of E. coli strain MG1655 [Δ(\(\upvarphi\)80-attB) IS 5.11::CmR-Ptac21-purA]

To construct E. coli strain MG1655 [Δ(\(\upvarphi\)80-attB) IS5.11::CmR-Ptac21-purA], the native purA gene was PCR amplified with primers 6–7 (see Additional file 1: Table S1 for details) using the MG1655 DNA as a template and was then cloned into a pMW119-P lac -lacI vector that had been treated with SmaI. Plasmid pMW119-P lac -lacI-purA was treated with BamHI and KpnI endonucleases, and the resulting purA-containing fragment was recloned into a pAH162-TcR-2Ter [34] integrative plasmid. The resulting plasmid, pAH162-TcR-2Ter-purA, was used for \(\upvarphi\)80-mediated integration of the promoter-less purA gene into the artificial \(\upvarphi\)80-attB site of the MG1655 [Δ(\(\upvarphi\)80-attB) IS 5.11::\(\upvarphi\)80-attB] strain constructed earlier [34]. Expression of the purA gene was activated by λRed-mediated insertion of the Ptac21 promoter region with two point mutations in the -35 region (ttgaca of P tac was replaced with tttgca) upstream of the gene using primers 8–9 (see Additional file 1: Table S1 for details). The obtained strain MG1655 [Δ(\(\upvarphi\)80-attB) IS5.11::CmR-Ptac21-purA] was used as the donor of a purA-carrier expression cassette, marked by an excisable CmR-marker, for P1-duction of the corresponding gene in the different strains of interest.

Construction of E. coli strain MG1655 [CmR-PL-purH]

To construct E. coli strain MG1655 [CmR-PL-purH], the upstream region of the purH gene, which is associated with purD in a single operon, was modified by replacement of the native regulatory region with the λPL promoter by λRed recombination as mentioned above. To construct the PCR fragment for λRed recombination with an excisable cat marker and flanking sequences homologous to the purHD regulatory region, we used primers 10 and 11 (see Additional file 1: Table S1 for details).

Construction of strain MG1655 [∆pitA::KmR]

To delete pitA in E. coli strain MG1655, the λRed integration method was used as described above. To construct the PCR fragment for λRed recombination with an excisable kan marker and flanking sequences homologous to the pitA gene, we used primers 12 and 13 (see Additional file 1: Table S1 for details) with the pMW118-KmR plasmid as a template. The chromosomal deletion of pitA was confirmed by PCR.

Construction of E. coli strain MG1655 [CmR-PLtac-pitA]

To construct E. coli strain MG1655 [CmR-PLtac-pitA], the upstream region of the pitA gene, was modified by replacement of the native regulatory region with the λPL tac promoter ggcggtg-ttgaca-attaatcatcggctcgtataatgtgtggaat [hybrid of λPL (ggcggtg) and λPtac (attaatcatcggctcgtataatgtgtggaat) promoters. λPtac promoter-35 sequence and λPL promoter -35 sequence, shared by the two promoters (ttgaca)]. To construct the PCR fragment for λRed recombination with an excisable cat marker and flanking sequences homologous to the pitA regulatory region, we used primers 14 and 15 (see Additional file 1: Table S1 for details).

Construction of strain MG1655 [ΔpurR::CmR ΔpurH::KmR]

To design an E. coli strain of MG1655 with double deletions of the purR and purH genes, MG1655 [ΔpurR::CmR] (Table 1) was used as the recipient for P1-duction of the ΔpurH::KmR cassette from the Keio collection [33]. The presence of two chromosomal modifications, ΔpurR::CmR and ΔpurH::KmR, was confirmed by PCR with primers 16–17 and 18–19, respectively (see Additional file 1: Table S1 for details).

Construction of the KF37-based strain with PhoBR-independent activation of Pho regulon genes

The sequence of phoBDBD, which encodes the C-terminal DNA-binding domain of PhoB [48] [specifically, (aa 1–3 of PhoB)-(Glu-Phe)-(aa 126–229 of PhoB)], was amplified by PCR using primers 20–21 (see Additional file 1: Table S1 for details) using the MG1655 chromosome as a template. Both the PCR product and integrative plasmid pAH162-TcR-2Ter (Table 1) were treated with SalI and SmaI restriction enzymes and then ligated with T4 DNA ligase. The ligation mixture was transformed into the CC118 λpir+ strain (Table 1), and the resulting pAH162-TcR-2Ter-phoBDBD plasmid, containing the promoter-less phoBDBD gene, was inserted into the yibH locus of the MG1655 [Δ(\(\upvarphi\)80-attB) ∆yibH::\(\upvarphi\)80-attB] chromosome by \(\upvarphi\)80-Int-mediated integration. The MG1655 [∆yibH::TcR-phoBDBD] strain was cured of the TcR marker using a pMWts-λInt/Xis helper plasmid. A PlacUV5 promoter with the KmR marker was inserted upstream of the phoBDBD gene using λRed recombination. The integrative DNA fragment was obtained by overlap extension PCR. To achieve this goal, the KmR marker was amplified from a pMW118-KmR template using primers 22–23 (see Additional file 1: Table S1 for details). Promoter PlacUV5 was amplified in parallel from a pML-PlacUV5-lacI [39] template using primers 24–25 (see Additional file 1: Table S1 for details). Extension PCR was performed by mixing the two initially amplified DNA fragments and using primers 22 and 26 (see Additional file 1: Table S1 for details). λRed recombination was applied to the MG1655 [∆yibH::phoBDBD] strain, and the resulting MG1655[∆yibH::KmR-PlacUV5-phoBDBD] strain was obtained. The strains MG1655-[∆yibH::TcR-phoBDBD] and MG1655 [∆yibH::KmR-PlacUV5-phoBDBD] were used as donors to individually transfer the constructs ∆yibH::TcR-phoBDBD and ∆yibH::KmR-PlacUV5-phoBDBD into strain KF37 by P1-duction.


AICAR controls its own formation in the His biosynthetic pathway

The enzymatic activity of HisG is subject to (i) feedback inhibition by His as the final pathway product, (ii) competitive inhibition by PR-ATP as a reaction product [6], and (iii) competitive inhibition by ADP and AMP, compounds that are structurally similar to its ATP substrate. The synthesis of each His molecule is accompanied by the release of one molecule of AICAR, an intermediate of de novo purine biosynthesis. In turn, AICAR is a well-known natural analog of AMP and can substitute for it in a number of biochemical reactions [49, 50]. Considering the structural similarity between AMP and AICAR (Fig. 2), not only AMP but also AICAR might negatively affect HisG enzymatic activity.
Fig. 2

Lineweaver–Burk plots. a Inhibition of HT-HisGWT by AMP. b Inhibition of HT-HisGWT by AICAR. Chemical structures of AMP (a) and AICAR (b)

To test this hypothesis, we measured the HisG activity of two purified His6-tagged enzymes, the wild type (HT-HisG) and a Fbr-mutant variant (HT-HisGE271K) that is resistant to feedback inhibition by His, in the presence of either AMP or AICAR (Fig. 2, Table 2). Figure 2 shows that the partial inhibition of HisG was detected for both tested compounds, but inhibition by AICAR was significantly more pronounced and was detected at low concentrations at which inhibition by AMP was nearly absent, that is in consistence with the literature data [51] (see Table 2). The dephosphorylated form of AICAR, (AICAr), does not affect HisG activity (data not shown). His biosynthesis is thus regulated not only by histidine itself as the terminal product of this pathway but also by AICAR as an AMP analog. Moreover, the biosynthesis of AICAR per se is under negative control via the initial step of His pathway, which is catalyzed by HisG.
Table 2

Specific activity of purified His6-tagged wild-type and feedback-resistant ATP-phosphoribosyltransferases and their inhibition by AMP and AICAR


ATP-phosphoribosyltransferase activity, µmol/min/mg

Inhibition, %





1 mM

20 mM

0.5 mM

1 mM

1 mM

20 mM

0.5 mM

1 mM


184 ± 9

195 ± 1

120 ± 15

151 ± 9

120 ± 5






120 ± 1

129 ± 12


67 ± 4

58 ± 8





Average data of 3 independent experiments are represented

nm not measured

AMP inhibits HisG activity due to competition with one of HisG’s native substrates, ATP [41]; AICAR could therefore be similar to AMP and thus exhibit an even stronger competitive inhibition of HisG.

Enhancement of AICAR conversion to ATP to intentionally overproduce His

The formation of AICAR by His biosynthetic enzymes is negatively controlled by AICAR inhibition of HisG. His production is thus directly regulated by this intermediate of purine biosynthesis, and a reduction in the AICAR pool due to its recycling to ATP may be one important way to overcome the bottleneck in His overproduction (Fig. 1). At a minimum, overexpressing purH and purA genes is necessary to enhance AICAR conversion to ATP (see Fig. 1). For purH overexpression, its native regulatory region in the chromosome of the MG1655 wild-type strain was replaced with the “strong” constitutive λPL promoter via λRed mediated recombination, and the MG1655 [CmR-PL-purH] strain was obtained (see Methods for details).

To increase purA gene expression, we specifically introduced an additional copy of the corresponding gene into the IS 5.11 locus of the MG1655 chromosome (as described in “Methods” section), followed by P1-duction of the CmR-Ptac21-purA cassette into several strains of interest. One of the obtained CmR-ductants, an MG1655-derivative that had earlier passed through several stages of selection for increased His overproduction, was used in the present study as the donor for P1-duction of the purA-carrier marked cassette. This cassette was P1-duced into the His-overproducing strain KF37 [13], followed by evaluation of His accumulation by the obtained CmR-ductants in test tube fermentations. Two groups of P1-ductants were selected according to their ability to overproduce His. The members of the first (main) group (more than 70% of the tested P1-ductants) increased His accumulation by approximately 20% over its levels in the initial KF37 strain. The members of the second (minor) group also exceeded KF37 in His overproduction, but the difference was approximately 6% (Table 3). Direct sequencing of the chromosomal regions flanking the bacterial locus IS5.11, the point of integration of the purA-carrier cassette in the P1-ductants obtained on the basis of KF37 and in the strain used as the donor for P1-duction, confirmed that the difference in His accumulation did not correspond to the structure of the cassettes, which were the same, in full agreement with the proposed design. The difference could be explained by the nonsense mutation that inactivated the pitA gene being closely coupled with the point of purA integration in the initial donor and in the tested P1-ductants that produced more His. This mutation was absent in the tested chromosomes of the P1-ductants from the second group. Moreover, the absence or presence of the mutation leading to pitA inactivation was confirmed by allele-specific PCR for the different members of the second and the first groups of P1-ductants, respectively (see Additional file 1: Table S1 for details, primers 1–3). One of the variants from the first group of P1-ductants that produced more His was cured of the CmR-marker due to Xis/Int-mediated recombination; the obtained markerless strain was assigned as EA79 and used for the following improvement process.
Table 3

Effect of PitA inactivation on His production by the strain KF37 [IS5.11::CmR-Ptac21-purA pitA]


pitA allele


His, g/L

His, %


pitA +

14.9 ± 0.4

3.3 ± 0.1


KF37 [IS5.11::CmR-Ptac21-purA, pitA+]

pitA +

16.1 ± 0.1

3.5 ± 0.1


KF37 [IS5.11::CmR-Ptac21-purA pitA]


15.6 ± 0.4

4.0 ± 0.1


Average data from 8 independent experiments are represented

Figure 3 shows that the overexpressed purH-carrier cassette was P1-duced from the strain MG1655 [CmR-PL-purH] into EA79 using CmR as a selective marker, and the marker was then excised to yield the EA83 strain, which accumulated 10% more His than EA79. In full accordance with the earlier supposition, the possible decrease in the intracellular AICAR pool due to the overexpression of purA and purH genes thus actually increased the level of His accumulation. At the same time, the occasional co-transduction of the mutation-inactivated pitA gene closely located to a P1-duced cassette had a much greater positive effect on His production than the overexpression of purA itself (see Table 3).
Fig. 3

Genealogy of His-producing strains with enhanced AICAR conversion into ATP. me marker elimination. Average data from 10 independent experiments are represented

PitA deficiency positively affects l-His production

To analyze the individual effect of PitA transporter elimination on His production, we deleted the corresponding gene in strain KF37, and the positive influence of PitA deficiency was confirmed (Table 4). In opposite, enhancement PitA transporter by introduction of the CmR-PLtac-pitA expression cassette, harboring overexpressed pitA gene under the strong λPLtac promoter control, into the His-producing strain KF37 lead to the drastically reduction of growth rate and His accumulation. The inactivation of pitB, which encodes a minor metal phosphate/H+ symporter, alone or in addition to the elimination of PitA did not influence the level of His accumulation (data not shown).
Table 4

Influence of PitA deficiency and enhancement on His production



His, g/L

His accumulation, %


14.9 ± 0.3

3.4 ± 0.1


KF37 [ΔpitA::KmR]

11.3 ± 0.5

3.8 ± 0.3


KF37 [CmR-PLtac-pitA]

7.1 ± 0.2

1.4 ± 0.2


Average data from 6 independent experiments are represented

The nature of this phenomenon is unclear, although several hypotheses concerning the positive influence of ΔpitA on His production were generated on the basis of the known properties of the PitA Me2+-Pi transporter. We supposed that the deletion of pitA can decrease import of Pi and Zn2+. The decreased import of Zn2+ may increase availability of Mg2+ for the efficient PPase-mediated PPi hydrolysis [24, 52] and result in the promotion of PPi-generating reactions, including those of the His biosynthetic pathway (Fig. 4, see “Discussion” for details). In turn, the His biosynthetic proteins HisG and HisI are PPi-releasing enzymes, and a metabolic significance of their function could thus be coupling with PPase activity to increase the intracellular Pi pool.
Fig. 4

Schematic view of the putative role of PitA deficiency in the cellular Pi/PPi balance and histidine biosynthetic pathway. ATP adenosine triphosphate, PP i inorganic pyrophosphate, P i inorganic orthophosphate, Me2+-P i metal phosphate complexes, PRPP phosphoribosyl pyrophosphate, PR-ATP phosphoribosyl-ATP, PR-AMP phosphoribosyl-adenosine monophosphate, ADP adenosine diphosphate, L-His l-histidine, HisG E271K feedback-resistant ATP-phosphoribosyltransferase, HisI phosphoribosyl-AMP cyclohydrolase/phosphoribosyl-ATP pyrophosphatase, Ppa inorganic pyrophosphatase, PitA low affinity Pi transport system, PstSCAB phosphate specific transport system

Influence of ΔpitA on Pho regulon induction and Pi uptake

To test the supposition concerning the decrease in inorganic phosphate import and probable alterations of the Pi pool in the absence of PitA, we evaluated the possible influence of PitA inactivation on Pho regulon gene activation. The activity of the bacterial alkaline phosphatase (EC, PhoA, the product of the phoA gene that belongs to the E. coli Pho regulon [5357]) was measured for this purpose. Growth in the MOPS-based minimal medium and Pi consumption were initially investigated for the wild-type strain MG1655, the His-producing strain KF37 and their respective pitA-deficient variants (see Additional file 3: Figure S2A and B). PhoA enzymatic activity was analyzed 24 h after Pi exhaustion. PitA transporter inactivation did not significantly change the growth efficiency and capability of Pi uptake of either MG1655 or KF37 under the tested conditions (see Additional file 3: Figure S2A), indicating that the total Pi uptake could be only slightly dependent on activity of PitA transporter. The insignificant effect of PitA transporter inactivation on capability of Pi uptake confirmed the knowledge concerning the main formation of intracellular Pi pools in E. coli. The variety of recent data show that the Pst system, but not PitA system, serves as the primary Pi transporter when Pi is in excess [22].

Interestingly, PhoA activity after Pi depletion was practically absent in the His-producing strain KF37, regardless of the pitA allele status and in contrast to the wild-type strain (Fig. 5). In the framework of the existing model of the control of Pho regulon [22, 58, 59], the hypothesis of possible AICAR negative influence on Pho regulon, in particular, PhoR kinase activity, was proposed. As known, E. coli PhoR histidine kinase has catalytic ATP-binding CA domain [58]. Pho regulon repression in case of His-producing strains could be explained by a reduction of PhoR’s ability to ATP-dependently autophosphorylate, which is necessary for the subsequent manifestation of its kinase activity in relation to PhoB when the Pho regulon activates (see Fig. 6, adapted from Hsieh and Wanner [22]). This reduction is probably based on an inhibition of ATP-dependent PhoR by AMP structural analog, AICAR, whose pool might be increased in His-producing strains.
Fig. 5

AP enzymatic activity under conditions of Pi limitation in His-producing and non-producing E. coli strains with different pitA alleles. MG1655, wild-type MG1655; MG1655 ∆pitA, MG1655 [ΔpitA::KmR]; KF37, MG1655+purR P his hisL′hisGE271KDCBHAFI]; KF37 ∆pitA, KF37 [ΔpitA:: KmR]. Average data from 3 independent experiments are represented, and error bars show the standard deviation (SD). PhoA enzymatic activity was analyzed 24 h after Pi exhaustion

Fig. 6

Proposed model of Pho regulon expression control. a Activation of the Pho regulon during Pi limitation. b Partial deactivation of the Pho regulon during Pi limitation in the presence of AICAR. PhoR histidine kinase, PhoB response regulator, PstSCAB phosphate-specific ABC transporter, PhoU negative regulator of PhoR, AICAR 5-aminoimidazole-4-carboxamide ribonucleotide, P i inorganic phosphate

Changes in the activity of PhoA in response to AICAR oversynthesis

To study whether the Pho regulon is repressed by AICAR, we constructed a strain with elevated intracellular levels of this metabolite and tested the activation of the Pho regulon in this strain under Pi-limiting conditions. Overproduction of AICAR was stimulated by inactivation of the PurR, a repressor of purine biosynthetic genes, and AICAR metabolism was simultaneously prevented by purH gene deletion. The AICAR pool was increased in the resulting double mutant strain MG1655 [ΔpurR::CmR ΔpurH::KmR], and this increase was detected by the appearance of AICAr, the dephosphorylated form of AICAR, in the culture broth (Table 5).
Table 5

Alkaline phosphatase (PhoA) activity under conditions of AICAR overproduction


AICAr, mg/L

PhoA activity, µmol/min/mg

MG1655 (wild-type)

< 0.1

410 ± 7

MG1655ΔpurR::CmR ΔpurH::KmR


261 ± 1

PhoA enzymatic activity was analyzed 2 h after Pi exhaustion. Average data of PhoA activity from 3 independent experiments are represented

We cultivated the E. coli wild-type strain and the strain overproducing AICAR under Pi-limiting conditions (their growth and Pi exhaustion kinetics are shown in Additional file 4: Figure S3). PhoA enzymatic activity was analyzed 2 h after Pi exhaustion. As expected, the level of PhoA alkaline phosphatase activity was appreciably less in the AICAR-overproducing strain than the wild-type strain (Table 5) but significantly higher than the KF37-based His-producing strains (compared in Fig. 5). These results collectively supported the hypothesis that an increased AICAR pool (in AICAR- and His-overproducing strains) was the main reason for the decreased expression of the Pho regulon genes and were likely due to AICAR inhibiting PhoR autophosphorylation, which in turn restricted the phosphorylation of PhoB.

Pi-independent activation of phoA gene expression in the His-producing strain KF37

The expression of a truncated PhoB regulator containing only the DNA-binding C-terminal domain (PhoBDBD [48, 57, 60]) was examined to confirm the normal functionality of the phoA gene in the KF37 His-producing strain. The expression of PhoBDBD provides a Pi-independent activation of the Pho regulon [48]. PhoA synthesis in the growing cells was easily visualized as blue-colored colonies on plates containing medium supplemented with the specific chromogenic substrate 5-bromo-4-chloro-3-indolyl phosphate (BCIP). Derivatives of the His-producing strain KF37 that also harbored the cassette containing the PhoBDBD gene under the control of the IPTG-inducible promoter PlacUV5 synthesized PhoA after induction, as did derivatives of the wild-type strain MG1655 harboring the same cassette (Fig. 7). This finding supports the hypothesis that His overproduction affects the function of the two-component signal transduction system PhoBR rather than the Pho regulon genes themselves.
Fig. 7

Strains expressing the phoBDBD gene. 1 KF37 [ΔyibH::TcR-phoBDBD], 2 KF37 [∆yibH::TcR-phoBDBD], 3 KF37 [ΔyibH::KmR-PlacUV5-phoBDBD]. 4 KF37 [ΔyibH::PlacUV5-phoBDBD], 5 KF37 [ΔyibH::PlacUV5-phoBDBD], 6 KF37, 7 MG1655 and 8 MG1655 [ΔyibH::PlacUV5-phoBDBD] after overnight incubation on LB agar supplemented with 50 mg/L BCIP without (a) and with (b) 500 µM IPTG addition


AICAR is formed during His biosynthesis and in the purine biosynthetic pathway [14, 29]. The negative effects of intracellular AICAR accumulation on different aspects of bacterial metabolism, such as adenosine homeostasis, gluconeogenesis, and thiamine synthesis, were investigated in S. enterica [61]. Moreover, several studies suggested that AICAR accumulation represses not only purine biosynthesis [62] but also one or more steps of the His pathway in S. cerevisiae [63]; however, the mechanism underlying such effects remains unclear. On the other hand, AICAR acts as a natural analog of AMP and can substitute for it in some enzymatic reactions. In turn, AMP inhibits enzymes involved in de novo purine and His biosynthetic pathways in which AICAR is formed as an intermediate [41]. We therefore hypothesized that AICAR controls its own formation, particularly in the His biosynthetic pathway. This supposition was investigated in this study by an examination of the possible influence of AICAR on the conversion of PRPP and ATP to PR-ATP, which is accomplished by HisG. The inhibitory effect of AICAR on HisG activity in vitro was demonstrated for the first time. The inhibition of HisG by high concentrations of AMP was also confirmed. Both tested substances compete with ATP in HisG-catalyzed reactions, suggesting that similar mechanisms are present for enzyme-ligand interactions. Figure 2 shows that AICAR was an even more effective inhibitor than AMP. AICAR thus controls its own formation with regard to the His biosynthetic pathway. Similar negative control may occur during AICAR formation via de novo purine biosynthesis; in this case, AICAR might control PurF enzyme activity, which is known to be negatively affected by AMP. In addition, some other possibilities for His biosynthesis regulation with participation of AICAR could be investigated. It is known that AICAR is identified as an alarmone that senses e.g. 10-formyl-tetrahydroflate deficiency in bacteria and activates a conserved riboswitch to regulate the expression of tetrahydrofolate genes involved in one-carbon metabolism [64, 65]. Therefore, it would be particularly interesting to study further the other aspects of the influence of AICAR on His biosynthesis, especially, considering close relationship of this biosynthetic pathway with one-carbon metabolism.

His overproduction cannot be engineered solely by enabling the resistance of the first biosynthetic enzyme, HisG, to feedback inhibition (see e.g., [13]). The effective metabolism of AICAR to ATP is also required to prevent the undesirable accumulation of AICAR, which is a HisG inhibitor, and to enhance the supply of ATP, which is an essential component for the synthesis of this amino acid. The effectiveness of such an approach was tested, yielding the E. coli His-overproducing strain EA83, which enhances AICAR regeneration into ATP (Fig. 3).

Furthermore, we found that a deficiency in PitA of the Me2+-Pi transport system considerably increased His accumulation (Table 3). A spontaneous mutation that inactivated the pitA gene was accidentally selected during the process of constructing the His-overproducing strains. The mechanism underlying this effect is not clear. Nevertheless, effective Zn2+-Pi import via the PitA transporter under conditions of excess extracellular phosphate and Zn2+ ions may restrict the total PPi hydrolysis efficacy, thereby inhibiting the His biosynthetic pathway reactions (Fig. 4). In E. coli, the hydrolysis of pyrophosphate (PPi) to Pi is catalyzes by inorganic pyrophosphatase (PPase, EC, the ppa gene product) in the presence of different divalent metal ions (e.g., Mg2+, Zn2+, Mn2+, and Co2+), but only using Mg2+ as physiological activator does PPase significantly increase the efficiency and specificity of PPi hydrolysis [52, 66, 67]. However, this specificity is lost when transition metal ions such as Zn2+, Mn2+ and Co2+ are used as cofactors, leading to markedly increased activity against polyphosphates [66] and the ability to hydrolyze organic tri- and diphosphates such as ATP and ADP [68, 69]. To test this hypothesis, we examine the effect of PitA deficiency on capability of Pi. Deletion of the pitA gene did not significantly decrease the import of Pi in the mutant strains from its levels in the PitA+ E. coli strains (Pi consumption values for both types of strains were within experimental uncertainty, see Additional file 3: Figure S2). These results correlate well with main formation of intracellular Pi pools by a basal level of Pst transporter activity in conditions of excess extracellular Pi [22]. The intracellular Pi pool was thus expected to be only slightly lower in pitA mutants than wild type. However, the deletion of pitA could additionally decrease import of Zn2+ ions and perhaps significantly increase the portion of PPase in complex with Mg2+, which improves enzyme–substrate specificity [70]. Decreasing the import of both Pi and Zn2+ could thus promote PPi-releasing reactions (in the His biosynthetic pathway, in particular) due to subsequent thermodynamically defensible hydrolysis of PPi to Pi.

To elucidate the roles of the other known Zn2+ transport systems during His production, we tested variants of a His-producing strain with a deleted znuA gene, which encodes the substrate-binding component of the ZnuABC high-affinity zinc uptake system [71], for His accumulation; the znuA deletion did not affect cell growth and had no positive effect on the rate of His accumulation (data not shown). These findings support the supposition that the effect of PitA deficiency on His production could not be explained solely by a decrease in Zn2+ uptake.

The last new feature that was detected for the His-producing strains in the present study was the significantly decreased Pho regulon activation in the growing cells after the depletion of the Pi that had been initially added in the cultural medium. This effect may have been based on the two-component PhoBR system and was likely to depend on AICAR-dependent inhibition of the PhoR autophosphorylation process. Indeed, the level of PhoA activation in the specially constructed AICAR-overproducing strain was lower than that of the wild-type strain, but this effect was not so pronounced as in His-producing cells.


Both the native HisG and feedback resistant HisGE271K from E. coli were shown to be sensitive to inhibition by AICAR. Accordingly, the production of l-histidine was improved in E. coli through the construction of strain EA83. In this strain, the AICAR regeneration pathway was enhanced via purA and purH gene overexpression and PitA-dependent phosphate/metal transport system was inactivated. Moreover, PhoA enzymatic activity was shown to be almost absent in the His-producing strain after Pi depletion in the medium, leading to the hypothesis that Pho regulon depression is associated with the inhibitory effect of AICAR, which is produced during the excessive production of His and acts as a regulatory molecule.





feedback resistance


phosphoribosyl pyrophosphate




5-aminoimidazole-4-carboxamide ribonucleotide, phosphorylated form, ZMP


5-aminoimidazole-4-carboxamide-1-beta-d-ribofuranosyl-5′-monophosphate, dephosphorylated form


5′-phosphoribosyl-5-formamido-4-imidazole carboxamide


inorganic phosphate


inorganic pyrophosphate


PstSCAB, phosphate specific transport


phosphate inorganic transport


metal phosphate (MeHPO4) complexes


thin layer chromatography






optical density


standard deviation


polymerase chain reaction


overlap extension PCR


sodium dodecyl sulfate polyacrylamide gel electrophoresis




alkaline phosphatase

Pho regulon: 

phosphate regulon


isopropyl β-d-1-thiogalactopyranoside


5-bromo-4-chloro-3-indolyl phosphate disodium salt


Authors’ contributions

EAM carried out the molecular genetic studies, constructed and analyzed the strain with improved His production ability, coordinated the work, and drafted the manuscript. IAB designed and performed the enzymatic study of HisG and participated in drafting the manuscript. AAK designed and performed the expression analysis of Pi-independent phoA gene activation. ABR participated in the strains and plasmids construction. SVM coordinated the work and helped to draft the manuscript. NVS designed and supervised the study and coordinated the writing of the manuscript. All authors read and approved the final manuscript.


We thank Dr. E.A. Kutukova and Dr. E.A. Slivinskaya for contributing to the construction of the strain with intracellular AICAR accumulation and PA Safronov for drawing the chemical structures in the figures.

Competing interests

The authors declare that they have no competing interests.

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Consent for publication

Not applicable.

Ethics approval and consent to participate

Not applicable.


Funding information is not applicable/no funding was received.

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Authors’ Affiliations

Ajinomoto-Genetika Research Institute, Moscow, Russian Federation


  1. Kopple JD, Swendseid ME. Evidence that histidine is an essential amino acid in normal and chronically uremic man. J Clin Invest. 1975;55(5):881–91.View ArticleGoogle Scholar
  2. Klyachko EV. Shakulov RS. Kozlov Yu I. Patent USA. 2008; No 7399618 B2.Google Scholar
  3. Ikeda M. Amino acid production processes. Adv Biochem Engin Biotechnol. 2003;76:1–36.Google Scholar
  4. Brenner M, Ames BN. Histidine regulation in Salmonella typhimurium. Metab Pathways. 1971;5:349–87.Google Scholar
  5. Alifano P, Fani R, Lio P, Lazcano A, Bazzicalupo M, Carlomagno MS, Bruni CB. Histidine biosynthetic pathway and genes: structure, regulation, and evolution. Microbiol Rev. 1996;60:44–69.Google Scholar
  6. Winkler ME, Ramos-Montanez S. Biosynthesis of histidine. EcoSal-Escherichia coli and Salmonella: cellular and molecular biology. Washington, DC: ASM Press; 2009.Google Scholar
  7. Morton DP, Parsons SM. Inhibition of ATP phosphoribosyltransferase by AMP and ADP in the absence and presence of histidine. Arch Biochem Biophys. 1977;181:643–8.View ArticleGoogle Scholar
  8. Martin RG. The one operon-one messenger theory of transcription. Cold Spring Harbor SympQuant Biol. 1963;28:357–61.View ArticleGoogle Scholar
  9. Hoppe I, Johnston HM, Biek D, Roth JR. A refined map of the hisG gene of Salmonella typhimurium. Genetics. 1979;92:17–26.Google Scholar
  10. Zhang Y, Shang X, Deng A, Chai X, Lai S, Zhang G, Wen T. Genetic and biochemical characterization of Corynebacterium glutamicum ATP-phosphoribosyltransferase and its three mutants resistant to feedback inhibition by histidine. Biochimie. 2012;94(3):829–38.View ArticleGoogle Scholar
  11. Kulis-Horn RK. Corynebacterium glutamicum ATP-phosphoribosyl transferases suitable for l-histidine production—strategies for the elimination of feedback inhibition. J Biotechnol. 2015;206:26–37.View ArticleGoogle Scholar
  12. Astvatsaturianz GV, Lisenkov AF, Smirnov IuV, Shakulov RS. Isolation of mutants with impaired retroinhibition of histidine biosynthesis. Genetika (Russian). 1988;24(11):1928–34.Google Scholar
  13. Doroshenko VG, Lobanov AO, Fedorina EA. The direct modification of Escherichia coli MG1655 to obtain histidine-producing mutants. Prikl Biokhim Mikrobiol. 2013;49(2):149–54.Google Scholar
  14. Bochner BR, Ames BN. ZTP (5-amino 4-imidazole carboxamide riboside 5′-triphosphate): a proposed alarmone for 10-formyl-tetrahydrofolate deficiency. Cell. 1982;29:929–37.View ArticleGoogle Scholar
  15. Zhang Y, Morar M, Ealick SE. Structural biology of the purine biosynthetic pathway. Cell Mol Life Sci. 2008;65(23):3699–724.View ArticleGoogle Scholar
  16. Honzatko RB, Fromm HJ. Structure-function studies of adenylosuccinate synthase from Escherichia coli. Arch Biochem Biophys. 1999;370(1):1–8.View ArticleGoogle Scholar
  17. Nath S. The molecular mechanism of ATP synthesis by F1F0-ATP synthase: a scrutiny of the major possibilities. Tools Appl Biochem Eng Sci. 2002;74:65–98.View ArticleGoogle Scholar
  18. Wanner BL. Phosphorus assimilation and control of the phosphate regulon. In: Neidhardt FC, editor. Escherichia coli and Salmonella: cellular and molecular biology. 2nd ed. Washington, DC: ASM Press; 1996. p. 1357–81.Google Scholar
  19. Rosenberg H, Gerdes RG, Chegwidden K. Two systems for the uptake of phosphate in Escherichia coli. J Bacteriol. 1977;131(2):505–11.Google Scholar
  20. Willsky GR, Malamy MH. Effect of arsenate on inorganic phosphate transport in Escherichia coli. J Bacteriol. 1980;144(1):366–74.Google Scholar
  21. Willsky GR, Malamy MH. Characterization of two genetically separable inorganic phosphate transport systems in Escherichia coli. J Bacteriol. 1980;144(1):356–65.Google Scholar
  22. Hsieh YJ, Wanner BL. Global regulation by the seven-component Pi signaling system. Curr Opin Microbiol. 2010;13:198–203.View ArticleGoogle Scholar
  23. van Veen HW, Abee T, Kortstee GJ, Konings WN, Zehnder AJ. Translocation of metal phosphate via the phosphate inorganic transport system of Escherichia coli. Biochemistry. 1994;33:1766–70.View ArticleGoogle Scholar
  24. Jackson RC, Binet MR, Lee LJ, Ma R, Graham AI, McLeod CW, Poole RK. Expression of the PitA phosphate/metal transporter of Escherichia coli is responsive to zinc and inorganic phosphate levels. FEMS Microbiol Lett. 2008;289:219–24.View ArticleGoogle Scholar
  25. Beard SJ, Hashim R, Wu G, Binet MR, Hughes MN, Poole RK. Evidence for the transport of zinc(II) ions via the pit inorganic phosphate transport system in Escherichia coli. FEMS Microb Lett. 2000;184:231–5.View ArticleGoogle Scholar
  26. Harris RM, Webb DC, Howitt SM, Cox GB. Characterization of PitA and PitB from Escherichia coli. J Bacteriol. 2001;183:5008–14.View ArticleGoogle Scholar
  27. Hayashi S, Koch JP, Lin EC. Active transport of l-α-glycerophosphate in Escherichia coli. J Biol Chem. 1964;239:3098–105.Google Scholar
  28. Pogell BM, Maity BR, Frumkin S, Shapiro S. Induction of an active transport system for glucose 6-phosphate in Escherichia coli. Arch Biochem Biophys. 1966;116:406–15.View ArticleGoogle Scholar
  29. Winkler HH. A hexose-phosphate transport system in Escherichia coli. Biochim Biophys Acta. 1966;117:231–40.View ArticleGoogle Scholar
  30. Sprague GF Jr, Bell RM, Cronan JE Jr. A mutant of Escherichia coli auxotrophic for organic phosphates: evidence for two defects in inorganic phosphate transport. Mol Gen Genet. 1975;143:71–7.View ArticleGoogle Scholar
  31. Sambrook J, Russell DW. Molecular cloning: a laboratory manual. Cold spring harbor: Cold spring harbor laboratory press; 2001.Google Scholar
  32. Studier FW, Moffatt BA. Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J Mol Biol. 1986;189(1):113–30.View ArticleGoogle Scholar
  33. Baba T, Ara T, Hasegawa M, Takai Y, Okumura Y, Baba M, Datsenko KA, Tomita M, Wanner BL, Mori H. Construction of Escherichia coli K-12 in-frame single-gene knockout mutants: the Keio collection. Mol Syst Biol. 2006;2006(2):0008.Google Scholar
  34. Minaeva NI, Gak ER, Zimenkov DV, Skorokhodova AY, Biryukova IV, Mashko SV. Dual-In/Out strategy for genes integration into bacterial chromosome: a novel approach to step-by-step construction of plasmid-less marker less recombinant E. coli strains with predesigned genome structure. BMC Biotechnol. 2008. Scholar
  35. Herrero M, de Lorenzo V, Timmis KN. Transposon vectors containing non-antibiotic resistance selection markers for cloning and stable chromosomal insertion of foreign genes in Gram-negative bacteria. J Bacteriol. 1990;172:6557–67.View ArticleGoogle Scholar
  36. Datsenko KA, Wanner BL. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc Natl Acad Sci USA. 2000;97(12):6640–5.View ArticleGoogle Scholar
  37. Haldimann A, Wanner BL. Conditional-replication integration excision and retrieval plasmid-host systems for gene structure–function studies of bacteria. J Bacteriol. 2001;183(21):6384–93.View ArticleGoogle Scholar
  38. Ublinskaya AA, Samsonov VV, Mashko SV, Stoynova NV. A PCR-free cloning method for the targeted phi80 Int-mediated integration of any long DNA fragment bracketed with meganuclease recognition sites into the Escherichia coli chromosome. J Microbiol Methods. 2012;89:167–73.View ArticleGoogle Scholar
  39. Skorokhodova AY, Katashkina ZI, Zimenkov DV, Smirnov SV, Gulevich AY, Biriukova IV, Mashko SV. Design and study on characteristics of auto- and smoothly regulated genetic element O3/PlacUV5/O lac lacI. Biotekhnologiya (Russian). 2006;5:3–21.Google Scholar
  40. Ames BN, Martin RG, Garry B. The first step of histidine biosynthesis. J Biol Chem. 1961;236:2019–26.Google Scholar
  41. Tébar AR, Ballesteros AO. Kinetic properties of ATP phosphoribosyltransferase of Escherichia coli. Mol Cell Biochem. 1976;11(3):131–6.View ArticleGoogle Scholar
  42. Waigh TA. The physics of living processes: a mesoscopic approach, vol. 21. New York: Wiley; 2014.Google Scholar
  43. Brickman E, Beckwith J. Analysis the regulation of Escherichia coli alkaline phosphatase synthesis using deletions and phi80 transducing phages. J Mol Biol. 1975;96(2):307–16.View ArticleGoogle Scholar
  44. Bradford MM. Rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem. 1996;72:248–54.View ArticleGoogle Scholar
  45. Ernster L, Lindberg O. Determination of organic phosphorus compounds by phosphate analysis. Methods Biochem Anal. 1956;3:1–22.Google Scholar
  46. Peterson GL. A simplified method for analysis of inorganic phosphate in the presence of interfering substances. Anal Biochem. 1978;84(1):164–72.View ArticleGoogle Scholar
  47. Moore SD. Assembling new Escherichia coli strains by P1-duction. Methods Mol Biol. 2011. Scholar
  48. Ellison DW, McCleary WR. The unphosphorylated receiver domain of PhoB silences the activity of its output domain. J Bacteriol. 2000;182:6592–7.View ArticleGoogle Scholar
  49. Hardie DG, Carling D, Carlson M. The AMP-activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryotic cell? Annu Rev Biochem. 1998;67:821–55.View ArticleGoogle Scholar
  50. Hardie DG, Salt IP, Hawley SA, Davies SP. AMP-activated protein kinase: an ultrasensitive system for monitoring cellular energy charge. Biochem J. 1999;338(3):717–22.View ArticleGoogle Scholar
  51. Dall-Larsen T. Stopped flow kinetic studies of adenosine triphosphate phosphoribosyl transferase, the first enzyme in the histidine biosynthesis of Escherichia coli. Int J Biochem. 1988;20(8):811–5.View ArticleGoogle Scholar
  52. Baykov AA, Cooperman BS, Goldman A, Lahti R. Cytoplasmic inorganic pyrophosphatase. Progr Mol Subcell Biol. 1999;23:127–50.View ArticleGoogle Scholar
  53. Torriani A. Influence of inorganic phosphate in the formation of phosphatases by Escherichia coli. Biochim Biophys Acta. 1960;38:460–9.View ArticleGoogle Scholar
  54. Heppel LA, Harkness DR, Hilmoe RJ. A study of the substrate specificity and other properties of the alkaline phosphatase of Escherichia coli. J Biol Chem. 1962;237:841–6.Google Scholar
  55. Malamy M, Horecker BL. The localization of alkaline phosphatase in E. coli K12. Biochem Biophys Res Commun. 1961;5:104–8.View ArticleGoogle Scholar
  56. Chang CN, Inouye H, Model P, Beckwith J. Processing of alkaline phosphatase precursor to the mature enzyme by an Escherichia coli inner membrane preparation. J Bacteriol. 1980;142(2):726–8.Google Scholar
  57. Blanco AG, Sola M, Gomis-Rüth FX, Coll M. Tandem DNA recognition by PhoB, a two-component signal transduction transcriptional activator. Structure. 2002;10:701–13.View ArticleGoogle Scholar
  58. Gardner SG, Miller JB, Dean T, Robinson T, Erickson M, Ridge PG, McCleary PW. Genetic analysis, structural modeling, and direct coupling analysis suggest a mechanism for phosphate signaling in Escherichia coli. BMC Genet. 2015. Scholar
  59. Carmany DO, Hollingsworth K, McCleary WR. Genetic and biochemical studies of phosphatase activity of PhoR. J Bacteriol. 2003;185(3):1112–5.View ArticleGoogle Scholar
  60. Bachhawat P, Swapna GVT, Montelione GT, Stock AM. Mechanism of activation for transcription factor PhoB suggested by different modes of dimerization in the inactive and active states. Structure. 2005;13:1353–63.View ArticleGoogle Scholar
  61. Bazurto JV, Heitman NJ. Downs Dm, aminoimidazole carboxamide ribotide exerts opposing effects on thiamine synthesis in Salmonella enteric. J Bacteriol. 2015;197(17):2821–30.View ArticleGoogle Scholar
  62. Rébora K, Daignan-Fornier B. Revisiting purine–histidine cross-pathway regulation in Saccharomyces cerevisiae: a central role for a small molecule. Genetics. 2005;170(1):61–70.View ArticleGoogle Scholar
  63. Hürlimann HC, Laloo B, Simon-Kayser B, Saint-Marc C, Coulpier F, Lemoine S, Daignan-Fornier B, Prinson B. Physiological and toxic effects of purine intermediate 5-amino-4-imidazolecarboxamide ribonucleotide (AICAR) in yeast. J Biol Chem. 2011;286(35):30994–1002.View ArticleGoogle Scholar
  64. Kim PB, Nelson JW, Breaker RR. An ancient riboswitch class in bacteria regulates purine biosynthesis and one-carbon metabolism. Mol Cell. 2015;57(2):317–28.View ArticleGoogle Scholar
  65. Ren A, Rajashankar KR, Patel DJ. Global RNA fold and molecular recognition for a pfl riboswitch bound to ZMP, a master regulator of one-carbon metabolism. Structure. 2015;23(8):1375–81. ArticleGoogle Scholar
  66. Höhne WE, Heitmann P. Tripolyphosphate as a substrate of the inorganic pyrophosphatase from baker’s yeast; the role of divalent metal ions. Acta Biol Med Germ. 1974;33:1–14.Google Scholar
  67. Josse J. Constitutive inorganic pyrophosphate of Escherichia coli. Purification and catalytic properties. J Biol Chem. 1966;231:1938–47.Google Scholar
  68. Schlesinger MJ, Coon MJ. Hydrolysis of nucleoside di- and triphosphates by crystalline preparations of yeast inorganic pyrophosphatase. Biochim Biophys Acta. 1960;41:30–6.View ArticleGoogle Scholar
  69. Kunitz M. Hydrolysis of adenosine triphosphate by crystalline yeast pyrophosphatase. J Gen Physiol. 1961;45:31–46.View ArticleGoogle Scholar
  70. Zyryanov AB, Shestakov AS, Lahti R, Baykov AA. Mechanism by which metal cofactors control substrate specificity in pyrophosphatase. Biochem J. 2002;367:901–6.View ArticleGoogle Scholar
  71. Patzer SI, Hantke K. The ZnuABC high-affinity zinc uptake system and its regulator Zur in Escherichia coli. Mol Microbiol. 1998;28(6):1199–210.View ArticleGoogle Scholar


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