Nonredundant Roles for Cytochrome c₂ and Two High-Potential Iron-Sulfur Proteins in the Photoferrotroph Rhodopseudomonas palustris TIE-1

R. palustris TIE-1 was grown in minimal medium as previously described (15), with the following modifications: the KH₂PO₄ concentration was reduced to 100 μM, and the NaHCO₃ concentration was increased to 30 mM. For hydrogen growth, the cooled medium was immediately aliquoted into sealed Balch tubes, and the headspace was flushed with 80% H₂–20% CO₂. Fifty-five to 60% of the vessel was left as headspace, which was pressurized to 5 to 10 lb/in². For acetate growth, 10 mM acetate was added to anaerobic medium with a headspace of 80% N₂–20% CO₂. Since the headspace was not required for the electron donor, the bottles were routinely filled almost to the top. For routine aerobic growth, YPS-MOPS growth medium (3 g/liter yeast extract, 3 g/liter peptone, 10 mM dibasic sodium succinate, 50 mM MOPS [morpholinepropanesulfonic acid], adjusted to pH 7 with NaOH) was used.

Culture growth was tracked by measuring the optical density at 650 nm (OD₆₅₀) of a 0.2-ml sample on a Synergy 4 plate reader (Biotech). Samples with an OD₆₅₀ of 0.3 were used for RNA extraction. Three milliliters of culture was added to 3 ml RNAlater (Ambion) in a Coy anaerobic chamber. Samples were centrifuged for 6 min at 6,000 × g and room temperature. The supernatant was poured off, and the pellets were frozen at −80°C until RNA extraction.

Cells were lysed using the lysis, digestion, and mechanical disruption protocol from Qiagen, and RNA purification was performed with a Qiagen RNeasy kit according to the manufacturer's instructions. Purified samples were treated with Ambion Turbo DNase using Ambion's rigorous treatment to remove contaminating DNA and quantified with a NanoDrop spectrophotometer. Forty nanograms of RNA was used to make cDNA using a Bio-Rad cDNA kit. One microliter of the cDNA reaction mixture was assayed in a 25-μl quantitative reverse transcription-PCR (qRT-PCR) reaction mixture using an iTAQ SYBR green Supermix with carboxy-X-rhodamine from Bio-Rad. Standard curves were made using RNA synthesized with an Ambion MEGAscript kit. The resulting RNA was diluted to a known concentration and treated in the same way as the experimental samples. qRT-PCR was performed with a 7500 fast real-time PCR system from Applied Biosystems. Data from four independent cultures were analyzed, and standard curves were generated using Sequence Detection software (version 1.4.0.25).

The strains used in this study are listed in Table 1, and the primers used in this study are provided in Table S1 in the supplemental material. To delete Rpal_4085, ∼1-kb fragments of the upstream and downstream regions were amplified using primer sets HipipUpF2/HipipUpR and HipipDownF/HipipDownR2. These fragments were recombined and cloned into pMQ84 using a Saccharomyces cerevisiae cloning system (16). The recombined fragment was cut out of pMQ84 and cloned into the suicide vector pJQ200SK. This vector was transformed into Escherichia coli S17-1 and mated into R. palustris TIE-1 as previously described (14). The strain in which pioC was replaced by Rpal_4085 was made in a similar fashion: the upstream, signal sequence, and downstream regions of PioC were amplified with the primer sets PioCmchF1C/PiC-HfusionsigR1b and PioC-HfusionsigF2/PioCHfusionR2, and the coding region (without the signal sequence) of Rpal_4085 was amplified with the primer set PioCHfusionF3/PioCmchR3C.

The cycA deletion strain was made as described above for the Rpal_4085 mutant using the primer sets Cyc2upF/Cyc2upR and Cyc2ddownF/Cyc2downR. The complementation strain was made in plasmid pBBR-MCS2 using the primers Cyc2CF and Cyc2CR. The PCR product and plasmid were cut with SpeI and HindIII. The ligation was performed with T4 ligase (New England Biolabs), and the resulting plasmid was transformed into E. coli DH5α electrocompetent cells. The successfully cloned plasmid was purified and sequenced to confirm that the full sequence was present. The plasmid was transformed into the ΔcycA strain and maintained with 100 μg/ml kanamycin.

The cell suspension assay for Fe(II) oxidation was performed as previously described (14). Briefly, cells grown on hydrogen were harvested by centrifugation when the OD₆₅₀ was ∼0.3, rinsed anaerobically in cell suspension buffer (50 mM HEPES, 20 mM NaCl, 20 mM NaHCO₃, pH 7), and resuspended in the same buffer plus ∼600 μM FeCl₂ to an OD₆₅₀ of ∼0.8. One hundred microliters of each sample was aliquoted into a clear flat-bottomed 96-well plate and incubated under a 60-W light bulb. Fifty microliters of samples was taken from a new well at each time point and combined with 50 μl ferrozine solution (0.1% [wt/vol] ferrozine in 50% ammonium sulfate). The absorbance was measured at 570 nm after a 10-min incubation, and the Fe(II) concentration was calculated on the basis of a standard curve.

The primers used in this study are listed in Table S1 in the supplemental material. Amplification of pioC was achieved by PCR with forward primer PioC-N-NcoCap and reverse primer PioC-C-XhoCap using genomic DNA extracted from aerobically grown R. palustris as the template. The PCR product was digested with the restriction enzymes NcoI and XhoI and ligated into the pET32h vector to generate plasmid pET32hPioC. PioC without its signal sequence was PCR amplified from pET32hPioC using forward and reverse primers PioCwithoutTAT and PioC-C-XhoCap, respectively. The resulting PCR product was cloned into pET32h by using NcoI and XhoI restriction enzymes.

The same expression protocol was used for the two proteins. PioC and Rpal_4085 were expressed in E. coli BL21 with a pET32h plasmid. The signal sequences predicted with TatP (17) were excluded from the resulting construct: thioredoxin–6× His–thrombin cleavage site–HiPIP. Both strains were grown in LB medium at 37°C. At an OD₆₀₀ of 1, expression was induced with 0.5 mM IPTG (isopropyl-β-d-thiogalactopyranoside) at 30°C. After 4 h, cells were harvested by centrifugation and resuspended in binding buffer (50 mM phosphate buffer, 300 mM NaCl, pH 8). Cells were lysed with a Thermo Scientific French press at 14,000 lb/in², and cell debris was removed by centrifugation. The resulting supernatant was loaded into a Qiagen Ni-nitrilotriacetic acid (NTA) agarose column equilibrated with binding buffer. Bound proteins were eluted with elution buffer (50 mM phosphate buffer, 300 mM NaCl, 250 mM imidazole, pH 8). After elution, the bound fraction was incubated with GE Healthcare thrombin overnight, according to the manufacturer's protocol. The enzymatic digest was dialyzed against binding buffer and reloaded into the reequilibrated Ni-NTA column. Target proteins did not bind to the column and were collected in the flowthrough. Purity was evaluated by SDS-PAGE with Coomassie blue staining. N-terminal sequencing showed that the obtained proteins were correctly digested by thrombin. Concentrations were estimated by UV-visible absorbance spectroscopy assuming a molar extinction coefficient of 16 mM⁻¹ cm⁻¹ at 396 nm. Nuclear magnetic resonance (NMR) data revealed a homogeneous population of correctly processed protein.

Cyclic voltammograms were recorded using an Autolab PSTAT10 potentiostat. The working electrode was a pyrolytic graphite-edge disc, a platinum wire was used as counter electrode, and an Ag/AgCl (1 M KCl) electrode was used as a reference. Reduction potentials are reported versus standard hydrogen electrode (SHE); experimental values were converted to potentials versus SHE by adding +222 mV to the readings made at 25°C. Prior to each experiment, the working electrode was washed with Millipore water (18 MΩ · cm). For each voltammogram, 20 μl of HiPIP at 5 μM in 50 mM Tris HCl, 300 mM NaCl, pH 9, buffer was used. The scan rate was 1 mV · s⁻¹. An initial potential of 750 mV was applied to the sample for 10 min before recording of each voltammogram. Data were processed with the SOAS program using a noise filter with an automatic threshold and subtraction of the background current (18).

The electron paramagnetic resonance (EPR) spectra of pure HiPIPs were recorded on a Bruker EMX spectrometer equipped with a dual-mode cavity and an Oxford Instruments continuous-flow cryostat. The experimental conditions were a temperature of 10 K, a microwave frequency of 9.66 GHz, microwave power of 6.346 mW, a modulation amplitude of 0.5 mT, and a receiver gain of 1.00 × 10⁵. The HiPIP concentration was 200 μM in 100 mM potassium phosphate buffer, 400 mM KCl, pH 8. Sample oxidation was achieved by addition of potassium ferricyanide at concentrations lower than the protein concentration, to avoid artifacts from unreacted ferricyanide.

Membrane fractions were prepared for EPR spectroscopy by two methods: for EPR spectra used in detection of natively expressed HiPIPs, the wild-type strain and the strain in which pioC was replaced by Rpal_4085 strains were grown on minimal medium with H₂-CO₂ to early stationary phase. Cell lysis was performed with the BugBuster reagent (EMD Millipore), according to the manufacturer's protocol, followed by ultracentrifugation at 200,000 × g. The pelleted membranes were resuspended in 20 mM phosphate buffer, pH 7, and oxidized with potassium hexachloroiridate(IV).

To prepare membranes for photooxidation experiments with pure proteins, the ΔpioC ΔRpal_4085 strain was grown anaerobically on minimal acetate medium to early stationary phase. The cell pellets were resuspended in 5 ml of 50 mM potassium phosphate buffer at pH 6 and passed three times through a Thermo Scientific French press at 25,000 lb/in². The cell lysate was centrifuged at 10,000 × g for 20 min to remove unbroken cells, and the supernatant was ultracentrifuged for 30 min in a Beckman-Coulter TLA-100.3 rotor at 60,000 rpm. The supernatant was discarded, and the pelleted membranes were resuspended in 4 ml of the same buffer. A total of 0.2 ml of 0.5 mM HiPIP solution was mixed with 0.1 ml of membrane suspension in a 3-mm EPR tube. The tubes were incubated for 30 min ∼10 cm from a 36-W white fluorescent lamp or in the dark, prior to freezing with liquid N₂.

The spectra of the membrane fractions and of the membrane suspension assay were recorded in a Bruker EMX spectrometer equipped with an ESR-900 continuous-flow helium cryostat. The experimental conditions were a temperature of 13 K, a microwave frequency of 9.39 GHz, microwave power of 2.012 mW, a modulation amplitude of 1 mT, and a receiver gain of 5.02 × 10⁴.

To determine the transcriptional response of Rpal_4085 to metal shocks, we grew 8-ml triplicate cultures anaerobically on minimal medium with acetate to mid-log phase and transferred them to an anoxic chamber. A 5-ml aliquot was removed and added to RNAlater as described above, and the cultures were shocked with either 2 μM CuCl₂, 50 μM NiCl₂, 1 mM CoCl₂, 1 mM FeCl₂, 1 mM MnCl₂, 1 mM ZnCl₂, 300 μM H₂O₂, or 1 mM CaCl₂. All metal stocks were prepared in anaerobic water. The concentrations were chosen to reflect the various levels of sensitivity to each compound that R. palustris growth displays. Shocked cultures were incubated under light for 20 min, and a second RNA sample was preserved.

Cytochrome c₂ was purified using a modified version of the native purification protocol described by Bartsch (19). Six liters of R. palustris was grown anaerobically on YPS-MOPS medium until stationary phase, harvested, washed in 1/10 volume of 100 mM Tris buffer, pH 8, and resuspended in the same buffer. Cells were lysed using a French press and ultracentrifuged for 1.5 h at 100,000 × g. The resulting lysate was fractionated by ammonium sulfate precipitation: ammonium sulfate solution was added to make a 30% solution, and the lysate was stirred for 10 min at 4°C and then centrifuged for 20 min at 25,000 × g. Additional ammonium sulfate was added to the supernatant to a final concentration of 90%. The lysate was again stirred and centrifuged, and the resulting pellet was resuspended in 10 mM Tris, pH 8, and desalted using a HiPrep 26/10 desalting column (GE Healthcare) into 1 mM Tris, pH 8.

The enriched cytochrome c₂ sample was further purified using ion-exchange chromatography. The desalted sample was passed through DEAE Sephacel resin (GE Healthcare) using a low-pressure Bio-Rad pump. The flowthrough, which contained cytochrome c₂, was adjusted to pH 5.5 by slow addition of 1 M acetic acid and then passed through CM Sepharose FastFlow resin equilibrated with 1 mM phosphate buffer, pH 6. The column was rinsed with 10 mM phosphate buffer, pH 6, and the protein was eluted with 20 mM Na-phosphate, pH 6. The eluate was diluted to 10 mM phosphate and loaded on a prepacked 1-ml CaptoS FastFlow column using an AKTA purifier fast-performance liquid chromatograph (GE). The column was rinsed with 10 mM Na-phosphate and eluted with 100 mM phosphate buffer, pH 6. The eluate was concentrated to 500 μl using a 3-kDa spin column and run on a gel filtration column as a final polishing step. The final protein was checked for purity by UV-visible spectra and run on an SDS-polyacrylamide gel stained with Coomassie.

Cytochrome c₂ from R. palustris TIE-1 was titrated in a Cary 5 UV/visible spectrophotometer using a home-built optical titration cell as described by Dutton (20). Cytochrome c₂ (100 μM) was titrated in the presence of the following redox mediators (in 50 mM MOPS at pH 7): ferrocene, p-benzoquinone, 2,5-dimethyl-p-benzoquinone, and 1,2-naphthoquinone (all at 20 μM) as well as 2,6-dichlorophenolindophenol at 5 μM. Ferricyanide and dithionite were added to, respectively, increase and lower the ambient midpoint potential.

Membrane fragments for reaction center reduction studies were prepared from acetate-grown cultures. Late-log-phase cells were lysed by 2 French press passages, spun at low speed to remove unbroken cells, and ultracentrifuged for 1 h at 150,000 × g. Membranes were rinsed once in 50 mM MOPS, pH 7, and resuspended in the same buffer.

Membrane suspensions were added to a round, stoppered quartz cuvette and measured on a Joliot type Jts-10 flash-induced spectrometer (Bio-Logic). Membrane-protein interactions were measured in the presence of ascorbate and myxothiazol to provide a reducing environment and inhibit the cytochrome bc₁ complex, respectively. Ascorbate was added from a 0.5 M stock solution. Myxothiazol was added from a 10 mM stock dissolved in ethanol and stored at −20°C. The reaction centers in the membrane fragments were photooxidized using a saturating flash from a custom-made (Daniel Beal, JBeamBio, France) light ring of actinic light-emitting diodes (800 nm), and changes in the absorbance at 435 nm (mainly originating from the photooxidized pigment) and 420 nm (dominated by redox changes of cytochrome c₂) were recorded. We chose to monitor cytochrome oxidation through changes at 420 nm because the change in alpha band absorption was obscured by electrochromic absorption changes in the same region. Spectral deconvolution was performed on the basis of the analysis of time-resolved flash-induced spectra, which will be described in a separate article (L. J. Bird, J. Lavergne, W. Nitschke, and D. K. Newman, unpublished data) focusing on whole-cell and in vitro analysis of light-induced electron transfer in R. palustris TIE-1.

Cytochrome c₂ is a common periplasmic electron shuttle between the cytochrome bc₁ complex and the reaction center; however, in some organisms, HiPIPs also assume this role (6). Given that TIE-1 has two HiPIPs as well as cytochrome c₂, we wanted to determine whether cytochrome c₂ is required for phototrophic growth. Accordingly, we made a clean deletion of the gene encoding cytochrome c₂ (cycA) and tested the resulting mutant strain for phototrophic growth with acetate or hydrogen as the electron donor. The ΔcycA mutant could not grow under these conditions, demonstrating that cytochrome c₂ is essential for photosynthesis in TIE-1 and that it cannot be replaced by either HiPIP (Fig. 1). The ΔcycA mutant also did not grow in the presence of Fe(II)-NTA; however, because TIE-1 growth on Fe(II) is unreliable when inoculated from an anaerobic culture, this result is not entirely conclusive.

We next sought to determine whether the HiPIPs PioC and Rpal_4085 could substitute for each other. Previous work had shown that although the TIE-1 ΔpioC mutant is unable to grow using Fe(II) as an electron donor, it still has some Fe(II) oxidation activity (14). We hypothesized that the residual Fe(II) oxidation activity might be due to the presence of another HiPIP (Rpal_4085) in the ΔpioC mutant, given that diverse proteins are known to donate electrons to the reaction center in other bacteria (21). To test this hypothesis, we deleted Rpal_4085 in both the wild-type and ΔpioC mutant backgrounds. The results were unexpected: the ΔpioC ΔRpal_4085 mutant was able to oxidize Fe(II) at the same rate as the ΔpioC mutant (Fig. 2), while the ΔRpal_4085 mutant grew on and oxidized Fe(II) at the same rate as the wild type. This suggests that Rpal_4085 does not substitute for PioC in the Fe(II) oxidation pathway. Given that both proteins are small redox-active proteins in the same family, we wondered why Rpal_4085 does not at least partially substitute for PioC in the iron oxidation pathway.

To determine whether Rpal_4085's inability to substitute for PioC was due to a lack of protein expression and/or a difference in localization between the two HiPIPs, we replaced the coding region of pioC with the coding region of Rpal_4085, leaving the twin arginine translocation (TAT) signal sequence of pioC intact. We used qRT-PCR to determine whether the mutant in which pioC was replaced by Rpal_4085 was properly expressing the Rpal_4085 gene. We found that in the wild-type strain grown on H₂, Rpal_4085 expression was significantly lower than pioC expression. In the mutant in which pioC was replaced by Rpal_4085, the expression of Rpal_4085 was increased to within the range of pioC expression in TIE-1 (Table 2). EPR spectroscopy of strains with a wild-type pioC operon and the substitution of Rpal_4085 for pioC indicated that HiPIP was present at very low levels in both samples (see Fig. S1 in the supplemental material). The resulting mutant (in which pioC was replaced by Rpal_4085) oxidized Fe(II) at the same rate as the ΔpioC mutant as shown in Fig. 2. This result strongly suggested that the Rpal_4085 protein cannot take on PioC's role in Fe(II) oxidation. This raised the following question: what is the biochemical basis of Rpal_4085's inability to substitute for PioC? As a step toward answering this question, we purified and characterized cytochrome c₂ and both HiPIPs.

We used EPR spectroscopy for two purposes: (i) to verify that both PioC and Rpal_4085 are typical HiPIPs and (ii) to determine whether they could be oxidized by the photosynthetic reaction center. We first expressed and purified the two HiPIPs from TIE-1 using a fusion with His-tagged thioredoxin. Because thioredoxin is placed at the N terminus, the periplasmic TAT signal sequence of the HiPIPs was removed in the cloning so the fusion protein would accumulate in the cytoplasm. This strategy, followed by affinity chromatography and proteolytic cleavage of thioredoxin, resulted in pure HiPIPs with a yield of 3 mg/liter culture. NMR experiments showed no evidence of significant amounts of apoprotein or of protein with an incorrectly assembled Fe-S cluster. EPR spectroscopy on pure protein confirmed that both PioC and Rpal_4085 displayed the characteristics of typical HiPIPs. This technique detects the presence of unpaired electrons which, in the case of metal clusters, have different spectral characteristics, depending on the nature of the cluster. Reduced HiPIPs have a diamagnetic ground state. Because only the ground state is occupied at the temperature of the EPR spectroscopy experiments, both proteins were EPR spectroscopy silent in the reduced form (not shown). In the oxidized form, the ground state is paramagnetic and HiPIPs display a characteristic EPR spectrum (22) (Fig. 3). The spectra in Fig. 3 show the characteristic signals of HiPIPs, where the main features have g values (proportionality constants, which reflect the environment of the electrons detected) of 2.12 and 2.03 for PioC and 2.12 and 2.04 for Rpal_4085. Both HiPIPs showed minor species, with the maximum g value being 2.08. These features are typical of HiPIPs and indicate that our purified proteins were correctly processed and active.

To test the ability of each HiPIP to donate electrons to the reaction center, we incubated a reduced sample of each protein with a suspension of membranes from the ΔpioC ΔRpal_4085 mutant either under light or in the dark. Because only oxidized HiPIPs show the characteristic EPR spectra, the detection of HiPIP by EPR spectroscopy in these experiments indicated that the protein could be oxidized by the reaction center. As observed through EPR spectroscopy (Fig. 4), PioC, but not Rpal_4085, is oxidized by the membrane suspension in a light-dependent manner.

We further characterized PioC and Rpal_4085 by measuring their reduction potentials. The reduction potentials were determined at pH 9 by cyclic voltammetry using a scan rate of 1 mV s⁻¹ (Fig. 5). Both proteins showed reversible reactions with midpoint potentials of +450 mV (PioC) and +470 mV (Rpal_4085). Faster scan rates resulted in irreversible reactions. At pH 7, scans were also irreversible, possibly due to poor interaction between the HiPIPs and the electrodes used.

The reduction potential of cytochrome c₂ was determined through optical redox titration, and cytochrome c₂ was found to have a midpoint potential of +350 mV.

Once we had confirmed the identities and determined the reduction potentials of the proteins, we further explored the interaction of the reaction center with cytochrome c₂ and PioC by using flash-induced absorbance spectrometry. We were able to monitor the absorbance changes caused by photooxidation and rereduction of membrane fragments in the presence of PioC or cytochrome c₂. In order to reduce all components prior to each experiment, ascorbate was added to the reaction mixture. Myxothiazol was used to inhibit the cytochrome bc₁ complex present in membrane fragments to ensure that cyclic electron flow did not take place, as this would have complicated the analysis. Because the EPR spectroscopy experiments with membranes indicated that Rpal_4085 could not reduce the reaction center at all, this experiment was not pursued for this protein.

A full description of the light-induced optical changes observed in TIE-1 will be published elsewhere (Bird et al., unpublished). For the present analysis, however, we used the relevant spectral parameters (peak wavelengths and extinction coefficients) obtained during this general spectroscopic and kinetic study to analyze the reactivities of cytochrome c₂ and PioC with the photosynthetic reaction centers (Fig. 6). Figures 6A and B compare the rereduction kinetics of the photooxidized primary donor (P⁺) in the reaction center in samples equilibrated with low, intermediate, and saturating concentrations of cytochrome c₂ and PioC, respectively, prior to the flash that triggered the reaction. These kinetics demonstrate that cytochrome c₂ and PioC differ significantly in their binding to and reaction with the reaction center.

Both PioC and cytochrome c₂ were able to reduce the photooxidized reaction center (Fig. 6A and B). Kinetic titrations with cytochrome c₂ showed an increase in the rate constant to a maximum of 200 s⁻¹ as the cytochrome concentration increased, indicating that the interaction between cytochrome c₂ and the reaction center is collisional (a pseudo-first-order reaction). The fact that only 50% of P⁺ was rereduced by cytochrome c₂ even at saturating concentrations of c₂ (Fig. 6A and B, inset) suggests that the membranes are present as equimolar populations of inside-out and right-side-out vesicles produced during the French press treatment. In contrast to cytochrome c₂, PioC's reduction of the reaction center had a time constant (k = 15 s⁻¹) that was independent of the concentration (Fig. 6B), indicating that it forms a complex with the reaction centers prior to the flash. The extent of PioC's donation to the reaction center, on the other hand, was concentration dependent (Fig. 6B, inset). Even at saturating concentrations of PioC, only roughly 60% of the accessible P⁺ (the population present in inside-out vesicles) was rereduced in a PioC-dependent manner. The supersaturating population of PioC therefore reduces only part of the photooxidized reaction center, demonstrating that PioC must be close to equipotential with the reduced/oxidized primary donor couple and that the observed kinetics correspond to achievement of redox equilibration rather than the intrinsic electron transfer rate, a parameter that can be observed only if a sufficient driving force (that is, a difference in redox midpoint potentials) for forward electron transfer is present. The strong binding of PioC to the reaction center was consistent with the characteristics of other HiPIPs (23), and redox equilibration between PioC and the reaction center was again consistent with the unusually high redox potential of PioC and the typically observed range of redox potentials for the primary donor pigment of the purple bacterial reaction (24).

Unlike PioC and cytochrome c₂, Rpal_4085 cannot be oxidized by the reaction center. We therefore looked for insight into its function by investigating its transcriptional response to metal stress. In a previously performed microarray (25), Rpal_4085 was upregulated by an anoxic Fe (II) shock, while PioC was not. We therefore hypothesized that it might be involved in mitigating metal toxicity. Using qRT-PCR, we tested the response of Rpal_4085 under anoxic conditions to the cations Mn(II), Fe(II), Co(II), Ni(II), Cu(II), Zn(II), Ca(II), and hydrogen peroxide (H₂O₂). With the exception of Zn(II) and Ca(II), these divalent metals are known to be redox active at physiological pH (19, 26–29). We used H₂O₂ as an oxidative stress-inducing control and Zn(II) and Ca(II) as nonredox active divalent metal controls. We found that in TIE-1 cultures grown on acetate, Rpal_4085 was highly upregulated in response to a shock with Mn(II), Fe(II), Co(II), and Ni(II) but not in response to a shock with Zn(II) or Ca(II) (see Fig. S2 in the supplemental material).