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The tricarballylate utilization locus (tcuRABC) of Salmonella enterica serovar Typhimurium is comprised of a 3-gene operon (tcuABC) that encodes functions that allow this bacterium to use tricarballylate as a source of carbon and energy, and the tcuR gene, which encodes a putative LysR-type transcriptional regulator. In our studies, transcription of the tcuABC operon peaked at mid-log phase, and declined moderately during stationary phase. This pattern was not due to a change in the amount of TcuR in the cell, as tcuR expression did not change under the conditions tested, and TcuR did not control tcuR expression. Tricarballylate was the co-inducer. tcuABC expression was negatively affected by the cAMP receptor protein (Crp). Expression of tcuABC was one order of magnitude higher in a crp mutant strain than in the crp+ strain; derepression of tcuABC expression was also observed in a strain lacking adenylate cyclase (Cya). At present, it is unclear whether the effect of Crp is direct or indirect. Studies with molecular mimics of tricarballylate showed that the co-inducer site restricts binding of structural mimics that contain a hydroxyl group. Two classes of TcuR constitutive variants were isolated. Class I variants responded to tricarballylate, while class II did not.
Tricarballylate is a citrate analog that is considered the causative agent of grass tetany, a ruminant disease characterized by an acute hypomagnesia . Tricarballylate is not catabolized by the ruminant or the rumen flora, hence it is excreted by the animal as a magnesium chelate . Unlike the normal rumen flora, Salmonella enterica serovar Typhimurium LT2 (hereafter referred to as S. enterica) can use tricarballylate as a carbon and energy source . The genome of S. enterica contains a 3-gene operon (tcuABC) dedicated to tricarballylate utilization (Fig. 4, supplemental material). The tcuABC operon contains all of the functions required for the catabolism of this tricarboxylic acid . The first gene of the operon (tcuA) encodes a protein with tricarballylate dehydrogenase activity, which converts tricarballyate to cis-aconitate, a Krebs cycle intermediate. TcuA is the only enzyme needed to catabolize tricarballylate; the tcuB and tcuC genes encode an electron transfer protein (required to re-oxidize the flavin cofactor of TcuA) and a tricarballylate transporter, respectively [23, 24].
Immediately upstream of the tcuABC operon is the tcuR gene, which encodes a putative LysR-type transcriptional regulator (LTTR)  (Fig. 4, supplemental material); tcuR is not part of the tcuABC operon. LTTRs are among the most ubiquitous forms of transcriptional regulators, and hundreds have been found in bacteria and some archaea . LTTRs have a distinct domain architecture (Fig. 5, supplemental information), with a helix-turn-helix in their N termini , while the C-terminal domains contain the co-inducer binding site and the oligomerization domain [1, 9, 11–13, 21, 22, 25, 35].
Here we provide in vivo evidence that the TcuR protein uses tricarballylate as its co-inducer for transcription of the tcuABC operon, and that the catabolite repression protein (Crp) negatively regulates expression of the operon. We also report the isolation and initial characterization of mutant alleles of tcuR that encode TcuR variants that activate tcuABC expression in the absence of tricarballylate.
A list of strains and plasmids used and their genotypes is provided in Table 4 (supplemental material). All chemicals were purchased from Sigma unless otherwise stated. Escherichia coli cultures were maintained in lysogenic (LB) broth [7, 8] (Difco). Nutrient broth (NB; Difco) was used as rich medium for S. enterica. Antibiotic concentrations were (in μg/ml): ampicillin (Ap), 100; chloramphenicol (Cm), 20; kanamycin (Km), 50; tetracycline (Tc), 20. No-Carbon E (NCE)  was used as minimal medium, and was supplemented with MgSO4 (1 mM), methionine (0.5 mM), and trace minerals [3, 15]. Whenever used as sole carbon and energy sources, citrate was at 20 mM, acetate at 30 mM, succinate at 30 mM, glucose at 10 mM, and cis-aconitate at 20 mM. When used in combination, succinate, acetate, and formate were all at 20 mM. All experiments were performed aerobically with shaking.
An in-frame deletion in tcuR was performed using a modification of the method described by Datsenko and Wanner . Briefly, the cat cassette of pKD3 was amplified using a 5′ primer containing 51-bp identical to the 5′ end of tcuR and a 3′ primer containing 51-bp identical to the 3′ end of tcuR. Manipulations were performed in strain JE6692 (Table 1). Insertion of the cat gene into tcuR was verified by DNA sequencing. The tcuR50::cat+ insertion was transduced into JE7212. Removal of the cat gene was performed as described .
S. enterica strain JE7212 (tcuA::MudJ(lacZ+ kan+) was mutagenized with N-methyl-N′-nitro-N-nitrosoguanidine (NG) as described by . Cells were incubated with NG for 40 min, and excess NG was removed by washing the cells with sterile phosphate buffer (0.1 M, pH 7.0) twice by centrifugation at 8000 g using a Beckman/Coulter Avanti J-25I microcentrifuge. Mutagenized cells were allowed to recover at 37°C until the culture reached an OD650 of ~1.5 before being plated for single colonies on NCE medium supplemented with lactose (10 mM). Large colonies were re-streaked on selective medium and saved. Mutant strains were reconstructed using phage P22 grown on the original mutant strains as donor, and strain TR6583 (tcuR+ tcuABC+) as recipient on NB agar plates containing kanamycin. Kmr transductants were replica-printed onto NCE medium supplemented with lactose. Lac+ colonies were freed of phage as described above. The tcuR gene from each Lac+ strain was sequenced to identify the mutations. Additionally, the first 500 bases immediately upstream of the tcuA start codon (tcuA promoter region) were sequenced, and no additional mutations were found in the tcuR mutant strains.
Unless otherwise stated, restriction and modification enzymes were purchased from MBI Fermentas and were used according to manufacturer’s instructions. All DNA manipulations were performed in E. coli DH5α/F′. Plasmids were transformed into E. coli cells by CaCl2 heat-shock . Plasmids isolated from E. coli were transformed into S. enterica via electroporation . Plasmid DNA was isolated using the Wizard Plus SV Plasmid Miniprep kit from Promega as per manufacturer’s instructions. DNA fragments were isolated from 1% (w/v) agarose gels and purified using the Qiaquick® gel extraction kit (Qiagen). PCR reactions were purified using the Qiaquick® PCR purification kit (Qiagen). Big-Dye® (ABI-PRISM) non-radioactive sequencing reactions were performed and the mixtures were resolved and analyzed at the Biotechnology Center of the University of Wisconsin-Madison.
Plasmids were propagated in E. coli strain DH5α/F′ except where noted. Genomic DNA for PCR was prepared from S. enterica strain JE6583 using the Wizard SV Genomic Purification System from Promega. All primers used for PCR amplifications were purchased from Integrated DNA Technologies.
The tcuC gene was sub-cloned from plasmid pTCU5  using 5′ SacI and 3′ HindIII sites and cloned into plasmid pBAD33  using the same restriction sites. Plasmid pTCU86 is 6.8-kb long and confers chloramphenicol resistance.
The 600-bp fragment immediately upstream of the ATG start site of tcuR was amplified using PCR primers containing a 5′ EcoRI site and 3′ BamHI site. The PCR product was cloned into plasmid pRS551  using the same restriction enzymes. Plasmid pTCU94 is13.1-kb long, and confers kanamycin and ampicillin resistance. pRS551 is a derivative of plasmid pBR322, which is an intermediate copy number vector (~20–30 copies per cell) .
β-Galactosidase activity was measured using established protocols . One unit of enzyme activity was defined as the amount of enzyme required to hydrolyze 1 nmol of o-nitrophenyl-β-D-galactopyranoside (ONPG) per min. Specific activity is reported as the number of units per OD650 unit. Enzyme activity was measured in mid-log cultures (i.e., OD650 ~0.4–0.6), unless otherwise noted. Optical density was monitored with a Spectronic 20D spectrometer. Unless otherwise noted, 5-ml cultures were used.
A reporter strain carrying both a tcuA::MudJ(lacZ+) transcriptional fusion and an in-frame deletion of tcuR (strain JE9315) was used to assess whether TcuR was required for tcuA expression. Strain JE9315 was transformed with plasmid pTCU5, which contained the wild-type allele of tcuC, the gene that encodes the tricarballylate transporter. The resulting strain, JE10458, was transformed with either plasmid pTCU26 (carries the wild-type tcuR+ allele) or a vector-only negative control. Cells were grown in NCE-glycerol (30 mM) ± tricarballylate (100 μM) to mid-log phase (OD650 ~0.4–0.5). Tricarballylate induced expression of tcuA::MudJ(lacZ+) by >22-fold when tcuR+ was provided in trans (Table 1; lines 2, 4). In contrast, we measured a 50-fold induction of tcuA expression in strain JE7212 (tcuA::MudJ(lacZ+), which carries a wild-type allele of tcuR on its chromosome, under the same growth conditions (NCE-glycerol + tricarballylate; Table 2, lines 5, 6). The observed differences in tricarballylate-induced tcuA expression likely reflect reduced expression of the transporter (tcuC) or the regulator (tcuR) genes. Support for this interpretation was obtained from experiments performed in the presence of high levels of arabinose (10 mM), which led to substantially higher induction of the tcuA-lacZ fusion by tricarballylate (111 ± 0.5 U/OD650). Lowering the arabinose concentration to 1 mM lowered tcuA-lacZ expression levels to 23.5 ± 3.0 U/OD650 when cells were grown with tricarballylate, versus 7.0 ± 0.3 when cells were grown in glycerol alone. Further lowering of the concentration of arabinose (0.5 mM) in the medium led to lower expression of the tcuA-lacZ fusion (9.6 ± 0.6 U/OD650) in medium containing tricarballylate, versus 5.8 ± 0.9 U/OD650 for glycerol alone.
Induction by tricarballylate depended on a functional TcuC protein. Expression of tcuA in strain JE7212 carrying the cloning vector (pBAD30) did not respond to tricarballylate (11 ± 3 U/OD650 in medium containing tricarballylate versus 4 ± 1 U/OD650 in glucose alone). In contrast, the same strain transformed with plasmid pTCU5 (tcuC+) responded to tricarballylate (807 ± 196 U/OD650 in medium containing tricarballylate versus 4 ± 1 U/OD650 in glucose alone). Thus, both TcuR and TcuC functions were required for tcuABC induction by tricarballylate.
We determined how much tricarballylate was required for maximal expression of the tcuABC operon. Strain JE7212 [tcuA33::MudJ(lacZ+)] was transformed with plasmid pTCU5 (tcuC+), the gene that encodes the tricarballylate transporter. A culture of the resulting strain (JE10614, Table 4, supplemental material) was grown to mid-log phase in NCE-glucose medium containing varying concentrations of tricarballylate, and β-galactosidase assays were performed. As shown in figure 1B, as little as 12 μM tricarballylate maximally induced tcuA-lacZ expression; clearly, the cell sensed and responded to extremely low levels of of tricarballylate in the medium. For all subsequent experiments, tricarballylate was added to 100 μM to ensure maximal tcuA-lacZ expression.
The observation that tricarballylate induced tcuA-lacZ expression in the absence of tricarballylate dehydrogenase (TcuA) function suggested that tricarballylate, not a catabolite of it, was the direct co-inducer of TcuR. To gain insights into the tricarballylate binding site of TcuR in the absence of a three-dimensional structure, strain JE7212 (tcuA33::MudJ(kan+)) was grown on different carbon sources (Table 2). Glucose, glycerol, citrate, isocitrate, and cis-aconitate failed to induce tcuA-lacZ expression. Lower but substantial levels of tcuA-lacZ expression were measured when succinate or acetate was used as carbon and energy source (Table 3). The effects of succinate and acetate depended on a functional TcuR protein, as strain JE9315 [ΔtcuR61 tcuA33::MudJ(kan+)] containing a deletion of tcuR failed to activate tcuA-lacZ on either succinate or acetate (3.0 ± 1.1 and 3.1 ± 1.8 U/OD650, respectively.) This result was consistent with succinate and acetate acting as co-inducers of TcuR. Because succinate is structurally similar to tricarballylate, (Fig. 3), we explained the stimulatory effect of succinate as molecular mimicry. To explore this idea further, we combined succinate with acetate and found that such a combination did not stimulate tcuA-lacZ expression relative to when succinate or acetate was used alone (Table 2). This result, while initially surprising, can possibly be explained if both succinate and acetate cannot simultaneously occupy the co-inducer site due to steric hindrance. Perhaps succinate and acetate displaced each other from the co-inducer site, thus resulting in poor activation of TcuR. To investigate the merit of this explanation we tested the combination of succinate and formate, which contains one less methyl group than acetate. The latter combination stimulated tcuABC to higher levels than those measured when succinate was used alone (Table 2), lending support to the idea that the methyl group of acetate sterically hinders the combined binding of succinate and acetate to the co-inducer site of TcuR. When used in combination with tricarballylate, neither succinate nor acetate abrogated the ability of tricarballylate to fully induce tcuA-lacZ expression (data not shown), which most likely reflects a higher affinity of TcuR for tricarballylate in comparison to these molecular mimics.
In contrast to the above findings, citrate and isocitrate were poor co-inducers of TcuR, suggesting that the co-inducer binding site of TcuR does not allow the presence of polar functional groups on the carbons connected to the carbonyl carbons of the co-inducer. One prediction for this hypothesis was that malate (hydroxysuccinate, Fig. 3) would be a poor inducer relative to succinate. Indeed, malate induced tcuA-lacZ expression only 17-fold compared to 50-fold for succinate (Table 2, lines 14, 9). Furthermore, the malate/formate combination increased tcuABC expression only 15-fold over un-induced levels. In contrast, for the succinate/formate combination we measured a 93-fold increased in expression, further supporting the idea that polar functional groups adjacent to the carbonyl groups may exert a negative effect on co-inducer binding. Further in vitro analysis of TcuR will address the true binding affinities of both succinate and acetate for TcuR as compared to tricarballylate.
Strain JE7212 was transformed with plasmid pTCU21, which carried the entire tcuABC wild-type operon; plasmid pTCU21 was previously shown to be able to complement the polar tcuA::MudJ(kan+) mutation . Plasmid pTCU21 contains 16 nucleotides immediately upstream of the tcuA start codon; the 16-bp region contains the tcuA Shine-Dalgarno sequence but lacks the tcuA promoter and putative TcuR binding sites. This construct allowed us to follow the temporal expression pattern of tcuABC expression while the cells are growing on tricarballylate. Notably, this construct also allowed us to maintain the proper stoichiometry of TcuR and TcuR-binding sites on the chromosome. To determine the temporal expression of the tcuABC operon, β-galactosidase assays were performed throughout the growth cycle when growing on tricarballylate (Fig. 1A). Expression of the tcuA-lacZ fusion peaked during exponential growth and dropped back to lag-phase levels when the cells began to enter stationary phase. These results suggested that the level of TcuR protein was not constant throughout the cell cycle, that proteins other than TcuR were involved in tcuABC operon regulation, or that a combination of these two possibilities occurred. We first tested whether TcuR regulated its own expression. Because expression of the tcuA-lacZ fusion peaked at mid-log phase, all experiments, including those shown in the preceding section above, were performed at mid-log phase to simplify comparisons.
The first 600 bp immediately upstream of the tcuR start codon were cloned into a promoter-less lacZ+ fusion vector (pRS551; ~20–30 copies per cell) to generate plasmid pTCU94. Expression of PtcuR-lacZ in strain TR6583 (tcuR+) remained constant in NB medium with or without 100 μM tricarballylate (1540 ± 20 versus 1450 ± 50 U/OD650, respectively). In addition, expression of PtcuR-lacZ did not change when tcuR was inactivated (JE7213; 1490 ± 40 U/OD650).)]. While most LTTRs repress their own transcription, those that do are most often divergently transcribed from the genes they regulate . Because tcuR is not divergently transcribed from tcuABC, the observation that neither TcuR nor tricarballylate had an effect on PtcuR-lacZtranscription was not surprising. We further tested whether the differences in growth rate expression of the tcuA-lacZ fusion were caused by differences in the level of TcuR throughout the growth phase. Expression of of PtcuR-lacZ (pTCU94) in the wild-type strain TR6583 was monitored at various time points when cells were growing on NCE-tricarballylate medium. The level of PtcuR-lacZ expression did not change throughout the growth curve (data not shown), suggesting that the increase in transcription of the chromosomal tcuA-lacZ fusion during exponential phase was not due to changes in the level of tcuR expression. However, we cannot at this time exclude the possibility that TcuR levels are regulated post-transcriptionally via targeted degradation or other means. In addition, there is evidence that pBR322 derivatives (i.e. pRS551) increase in copy number during stationary phase . Thus, it is possible that a decrease in PtcuR-lacZ is masked by the increase in copy number. Further in vitro characterization of TcuR should establish whether TcuR can bind to its own promoter and repress transcription.
We assessed whether the tcuABC operon was controlled by catabolite repression. For this purpose, we tested the effects of mutations in either adenylate cyclase (Cya) or cAMP receptor protein (Crp). Tricarballylate-dependent expression of the chromosomal tcuA-lacZ fusion in a strain of S. enterica carrying a null allele of the crp gene [JE7830 tcuA33::MudJ(lacZ+) crp::Tn10(tet+)] was >10-fold higher than the level of tcuA-lacZ expression in the crp+ strain (JE7212 tcuA33::MudJ(lacZ+), Fig. 2). Consistent with the involvement of the Crp/cAMP complex, strain JE10114 [tcuA33::MudJ(lacZ+) cya::Tn10(tet+)] carrying a null allele of adenylate cyclase (cya) also showed a substantial increase (6-fold) in tcuA-lacZ expression relative to the level measured in the cya+ strain. Addition of cAMP (1 mM) to the medium reduced tcuA-lacZ expression in the presence of tricarballylate >3.5-fold, and the introduction of plasmid pHY26-1, which carried an allele of crp that encoded a cAMP-independent Crp protein, reduced tcuA-lacZ expression ~2-fold. Reasons that pHY26-1 resulted in lower repression compared to the addition of cAMP may include improper gene dosage and/or partial activity of the constitutive Crp variant protein.
If Crp were acting as a repressor of the tcuABC operon, we would expect tcuA-lacZ expression to be activated during growth on glucose. Indeed, tricarballylate-induced tcuA-lacZ expression was higher in glucose-containing medium than either citrate or glycerol-containing medium (Table 2; lines 2, 4, 6). These results suggested that the Crp protein acted as a repressor of tcuABC expression. This was somewhat surprising, as Crp most commonly acts as an activator and only a few enzyme-encoding genes are repressed by Crp/cAMP . It is not obvious why the cell would use Crp to repress tcuABC expression. We speculate that the involvement of Crp in tcuABC expression reflects conditions where S. enterica encounters tricarballylate in the environment. The extant regulatory circuit of tcuABC expression allows S. enterica to catabolize tricarballylate in the presence of glucose. If glucose repressed tricarballylate catabolism in a canonical catabolite repression fashion, any tricarballylate that entered the cell would inhibit aconitase, preventing the synthesis of essential amino acid precursors such as α-ketoglutarate, succinyl-CoA and oxaloacetate.
It is possible that Crp acts as a repressor of tcuR and that Crp repression of tcuA-lacZ expression is at least partially the result of Crp-mediated repression of tcuR. We tested whether PtcuR-lacZ expression from plasmid pTCU94 differed in either a crp or cya background (strains JE10943 and JE10944, respectively) when the cells were grown in LB medium containing tricarballylate (100 μM). We measured no significant difference (1.2 fold) in PtcuR-lacZ expression in the crp or cya strains compared to that of the wild-type strain TR6583 (1600 ± 70 and 1790 ± 40 versus 1450 ± 70 U/OD650, respectively). Addition of cAMP (1 mM) to the medium reduced PtcuR-lacZ in the cya strain by 1.3-fold (1790 ± 40 versus 1360 ± 70 U/OD650, respectively). Furthermore, there was slight evidence of glucose activation, as expression of PtcuR-lacZ in the wild-type strain was only 1.1-fold increased for cells growing on glucose compared to LB (1620 ± 50 vesus 1450 ± 70 U/OD650, respectively). These results suggested that the Crp protein did not have an effect on tcuR expression. While it is possible that Crp could have affected the copy number of pTCU94, we found no evidence that this may be the case in the literature. Additionally, there are no notable consensus motifs for Crp-binding sites in the tcuR promoter region (see below). Direct in vitro binding studies between Crp and the tcuR promoter region would firmly establish whether Crp directly interacts with the tcuR promoter.
Crp-dependent repression of tcuA-lacZ expression and the increase in the expression of tcuA-lacZ during mid-log growth suggest that tcuABC regulation is complex. At present, it is unclear whether the effects of Crp are direct or indirect, and whether any other global regulators also control either tcuR or tcuABC operon expression. Noteworthy, there are no notable consensus motifs for Crp-binding sites in either the tcuR or tcuA promoter regions. If the Crp effect were indirect, we would hypothesize that Crp would act as an activator of a protein that represses tcuABC expression. Hence in the absence of Crp, and consequently of the putative repressor, tcuABC expression would increase. In vitro biochemical studies are needed to determine whether the effect of Crp on tcuABC expression is direct.
Chemical mutagenesis with NG was performed on strain JE7212 (tcuA33::MudJ(lacZ+) to isolate constitutive mutations in TcuR. We selected derivatives of strain JE7212 that formed large colonies on NCE-lactose medium lacking tricarballylate. Each mutant was reconstructed as described above, and in every case the Lac+ phenotype was co-transducible with the kan+ marker. We identified eight mutant tcuR alleles upon sequencing (Fig. 5, supplemental material). The promoter region of each tcuR strain was also sequenced, and no additional mutations were found. Several tcuR mutations were isolated between residues 241–244, the region of the protein containing the putative C-terminal multimerization domain. While the screen was likely non-saturating, several mutations were independently isolated twice (E42G, L264R) or thrice (A242T), highlighting the relevance of the position. Changes in the residues within the proposed co-inducer-binding domain were not isolated. Reasons why we did not find mutations in this domain could include the mutational bias of NG, the stringency of the screen and the need for multiple mutations to observe the desired phenotype.
Expression of the tcuA-lacZ fusion in the mutant strains was assessed during growth in rich medium with or without tricarballylate. While every mutant strain had at least a 25-fold increase in tcuA-lacZ expression in the absence of tricarballylate, there were two distinct phenotypes in the presence of tricarballylate. Class I mutations mapped within four residues of each other, and were defined as those alleles encoding TcuR variants whose activity was >2-fold higher when tricarballylate was added to the medium (Table 3). Class I mutations may bias TcuR towards achieving an active conformation in the absence of tricarballylate, but still allow a response to tricarballylate. Class II mutations, with the lone exception of S241C, were located elsewhere in TcuR, and responded poorly to tricarballylate (<2-fold induction of tcuABC). Class II mutations may enhance DNA-binding independently of the normally required conformational change. In fact, the E42G substitution (a class II mutation) falls within the putative helix-turn-helix, which would support this explanation. The L264R substitution (also class II) may also increase DNA-binding affinity, as substitutions near this residue in both NahR (regulator of naphthalene catabolism in Pseudomonas putida) and AmpR (regulator of β-lactamase expression in Citrobacter freundii) led to loss of DNA-binding in previous studies [5, 30].
Substitution G241C may be an outlier, as it falls within the group of residues with a Class I phenotype. In this case, it is possible that this TcuRG241C variant is locked in a conformation that prevents it from responding to tricarballylate. In vitro approaches are needed to analyze Class I and Class II mutations to determine their effects on DNA-binding and transcriptional activation.
This study establishes the in vivo basis for the regulation of the tcuABC operon by TcuR and its co-inducer, tricarballylate. We have shown that tricarballylate alone is sufficient for TcuR-mediated induction of tcuABC. Our results also demonstrate a role for the global regulator, Crp, in tricarballylate catabolism. The results described here should be valuable for future structure/function analyses of the important LTTR family. The response of TcuR to molecular mimics of tricarballylate including succinate and acetate may lead to increased understanding of how LTTRs recognize their co-inducers. In addition, the two classes of constitutively active mutants may lead to understanding of how LTTRs adopt a transcriptionally competent conformation. Current efforts are focused on obtaining a crystal structure of TcuR and performing detailed in vitro analysis.
This work was supported by PHS grant R01-GM62203 to J.C.E.-S. L.W.S. was the recipient of the NIH Undergraduate Scholarship. J.A.L. was supported in part by the Department of Bacteriology Jerome Stefaniak Predoctoral Fellowship and by the Molecular Biosciences Training Grant T32 GM07215. We thank Kathy Krasny for technical assistance.
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