Published: December 17th, 2018
Here, we present a protocol to use an anaerobic whole-cell microbial biosensor to evaluate how different environmental variables affect the bioavailability of Hg and Cd to bacteria in anoxic environments.
Mercury (Hg) bioavailability to microbes is a key step to toxic Hg biomagnification in food webs. Cadmium (Cd) transformations and bioavailability to bacteria control the amount that will accumulate in staple food crops. The bioavailability of these metals is dependent on their speciation in solution, but more particularly under anoxic conditions where Hg is methylated to toxic monomethylmercury (MeHg) and Cd persists in the rhizosphere. Whole-cell microbial biosensors give a quantifiable signal when a metal enters the cytosol and therefore are useful tools to assess metal bioavailability. Unfortunately, most biosensing efforts have so far been constrained to oxic environments due to the limited ability of existing reporter proteins to function in the absence of oxygen. In this study, we present our effort to develop and optimize a whole-cell biosensor assay capable of functioning anaerobically that can detect metals under anoxic condition in quasi-real time and within hours. We describe how the biosensor can help assess how chemical variables relevant to the environmental cycling of metals affect their bioavailability. The following protocol includes methods to (1) prepare Hg and Cd standards under anoxic conditions, (2) prepare the biosensor in the absence of oxygen, (3) design and execute an experiment to determine how a series of variable affects Hg or Cd bioavailability, and (4) to quantify and interpret biosensor data. We show the linear ranges of the biosensors and provide examples showing the method's ability to distinguish between metal bioavailability and toxicity by utilizing both metal-inducible and constitutive strains.
Mercury (Hg) is a global pollutant and its bioavailability to Hg methylating microbes is the first step towards its biomagnification through food webs and its possible neurotoxic effects in human and wildlife1. It is currently thought that microbial Hg methylation is an intracellular process that requires: i) the species of Hg to be bioavailable2,3,4,5,6,7 and ii) for the cell to be physiologically capable of methylating Hg8,9,10. Cadmium (Cd) bioaccumulates in organisms, but does not biomagnifies in foodwebs and is widely used in industrial and commercial processes that commonly cause acute exposures in people and the environment11. Although microbes play several key roles in the fate of Hg in the environment, most studies on Cd geochemistry and ecotoxicology focus on microbial eukaryotes12. Consumption of agricultural crops (e.g., rice) is the main source of direct exposure to cadmium; in this case, the bioavailability of Cd to microbes of the rhizosphere directly influences the amount plants can accumulate13.
Hg transport pathways are complex and possibly involve an active transport step14. When a transporter is involved, recent work suggested that HgII uses a ZnII or MnII transporter7,15,16,17. Whereas CdII is hypothesized to be accidentally transported into the cytosol through divalent metal transport pathways (particularly MnII or ZnII), mechanisms of CdII transport inside the cells remain speculative, and no Cd-specific transport pathway has been identified13,18. Regardless of the nature of the transporters involved, three mechanistic factors ultimately determine the ability of metals to enter a cell: i) the metal speciation in solution2,6,15,16,17,19,20,21,22,23, ii) the biophysiochemical properties of the cell membrane17,24,25,26,27,28,29,30,31, and iii) the ability for the metal to access a transport site7,32. Cd and Hg are unlikely to exists as free ions under microbial physiologically relevant conditions due to their high affinity for Dissolved Organic Matter (DOM), chelating contaminants (e.g., EDTA), or reduced sulfur moieties33,34,35 (CdII can exist as a free ion or form ion-pairs in the absence of these ligands). There is a lack of efficient methods in determining how these metal species are bioavailable under conditions relevant to their fate in the environment. For instance, Hg is methylated under anaerobic conditions14, and both cadmium and Hg are soft metals (or class B cations), requiring that their speciation be investigated under conditions that maintain the integrity of reduced sulfur groups.
Microbial biosensors are bacterial cells that emit a quantifiable signal in response to the intracellular concentrations of a metal, in this case Hg or Cd. As such, they are ideal tools to understand how metals enter a cell36, provided that exposure conditions are carefully controlled for. Hg biosensors typically contain gene fusions between the regulatory circuitry of the mer-operon (including genes encoding for the transcription regulator MerRas well as the operator and promoter regions of the operon), and reporting genes (e.g., lux, gfp, lac genes). When mercury is present in the cytoplasm, it will bind to MerR, resulting in transcription of the reporting genes and subsequent signal production19,37. Specific Cd biosensors are usually designed using the cadC, cadAC, zntA or zntR encoded transcription regulators38, but it is worth noting that MerR has a lower, yet quantifiable affinity to Cd5. Due to aerobic restriction of most commonly used luminescent or fluorescent reporter proteins, until recently microbial biosensors remained unable to offer insights into the biotransformation of metals under anoxic conditions. This makes anaerobic detection of metals bioavailability very difficult over a range of conditions relevant to their environmental fate, specifically in the presence of redox sensitive ligands (e.g., sulfide and thiols)4,5,39.
To alleviate the methodological hurdle of live imaging in the absence of oxygen, Drepper et al. (2007) have developed a flavin-based fluorescent protein (FbFp), based on light oxygen voltage domain of SB2 protein from P. putida. This protein family is able to fluoresce in the absence of oxygen40. Building on the work of Drepper et al., our lab designed an anaerobic biosensor allowing for the study of Hg bioavailability under oxic and anoxic conditions and over a wide range of salinity 17. In the current paper, we describe how to prepare and use the biosensor to test environmental variables' influence on Hg or Cd bioavailability. Although we developed the biosensor for HgII, we chose to perform experiments with CdII as a means to draw the reader's attention to the fact that biosensors may also respond to multiple stressors that are likely to co-occur in environmental matrices; in this case CdII was investigated because it is known to bind to the transcriptional regulator MerR5. Here, we show representative calibration and functional linear ranges with respect to either metal. We also give an example when the biosensor's results are conclusive (MgII and MnII on Hg bioavailability) and inconclusive (ZnII on Hg bioavailability).
1. Growth Media and Exposure Media Preparation
2. Preparation of Mercury and Cadmium Standards.
3. Preparation of the Biosensor for Anaerobic Exposure Assay
4. Exposure Assay
Figure 1: A 96 well plate (left) and a corresponding 4 x 8 grid containing PTFE vials (right) to be transferred to the plate. Please click here to view a larger version of this figure.
Note: When testing for the role of a variable on Hg uptake with the mercury inducible biosensor; two treatments are required for each variable: the treatment (biosensor + Hg + variable + nitrate) and its treatment blank (biosensor + variable + nitrate). When testing for the role of a variable on the physiology of the cell using the constitutive biosensor, two treatments are required for each variable: the treatment (biosensor + Hg + variable + nitrate) and treatment blank (biosensor + variable + Hg). Mercury may be replaced with Cadmium. Hg or Cd will become the variable when performing a calibration curve. The constitutive and inducible biosensors do not need to be run at the same time (in the same plate layout). A template example for the plate layout and corresponding 4 x 8 grid when testing a concentration range of magnesium (variable) is provided in Table 1).
|Hg induced biosensor + Hg + Nitrate + 0 mM Mg
|Hg induced biosensor + Nitrate + 0 mM Mg
|Constitutive biosensor + Hg + Nitrate + 0 mM Mg
|Constitutive biosensor + Hg + 0 mM Mg
|Hg induced biosensor+ Hg + Nitrate + 0.1 mM Mg
|Hg induced biosensor + Nitrate + 0.1 mM Mg
|Constitutive biosensor+ Hg + Nitrate + 0.1 mM Mg
|Constitutive biosensor + Hg + 0.1 mM Mg
|Hg induced biosensor + Hg + Nitrate + 1 mM Mg
|Hg induced biosensor + Nitrate + 1 mM Mg
|Constitutive biosensor + Hg + Nitrate + 1 mM Mg
|Constitutive biosensor + Hg + 1 mM Mg
|Hg induced biosensor + Hg + Nitrate + 10 mM Mg
|Hg induced biosensor + Nitrate + 10 mM Mg
|Constitutive biosensor + Hg + Nitrate + 10 mM Mg
|Constitutive biosensor + Hg + 10 mM Mg
Table 1: An example plate layout for using the biosensor to test Hg bioavailability (5 nM) over a gradient of Magnesium (0-10 mM)
5. Quantifying the Data
Fluorescence (t) = average(Tr1(t) – Tr1(t0), Tr2(t) – Tr2(t0), Tr3(t) – Tr3(t0)) – average(TBr1(t) – TBr1(t0), TBr2(t) – TBr2(t0), TBr3(t) – TBr3(t0)) (1)
Note: This should be made as a spreadsheet function. Proper propagation of error should also be calculated for each time point.
Figure 2: Corrected fluorescence data as a function of time. Fluorescence measured as relative fluorescence units (RFU) emitted by E. coli NEB5α harboring the pUC57merR-Pp (Inducible strain) over time with the addition of HgII (0-12.5 nM) under anaerobic conditions. Fluorescence was the average of 3 technical replicates at 37 ˚C. Please click here to view a larger version of this figure.
Note: There is no 0 nM Hg value on the graph, and all other Hg concentrations have been blanked to the 0 nM Hg as a treatment blank. Therefore, 0 nM Hg represents the x axis and any positive fluorescence represents fluorescence from Hg given any variable. It is optional to not blank the fluorescence in this manner, but the fluorescent curves will give misleading fluorescence curves if the variable tested has background fluorescence (i.e., dissolved organic matter itself will fluoresce and if there is no treatment blank containing just cells and the dissolved organic matter, increasing dissolved organic matter concentration will increase the fluorescent signal).
Once the fluorescence peaks have been quantified according to step 5.3, the result of the fluorescence peaks can be graphed according to the variable concentration illustrating how that variable affects the relative bioavailability of either Hg or Cd. For example, the calibration curve of fluorescence over [HgII] from Figure 2 will yield the inducible data presented in Figure 3A. For HgII calibration, the curve will always contain 3 components for the Hg-inducible strain; a threshold response of about 1-2 nM HgII before fluorescence signal production is linearly proportional to [HgII], the linear range where 5 nM HgII will reliably always be in the center of that range, and a plateau where increasing [HgII] will no longer increase fluorescence signal. No change in signal production on the constitutive strain shows that toxicity from [HgII] does not affect signal production. For CdII in Figure 3B, there are always 2 components for the inducible strain; a linear range where 200-300 nM CdII will reliably always be in the center of the linear range and a plateau. A decrease in fluorescence signal with increasing [CdII] shows that higher Cd concentrations are toxic to the cells and can explain a decrease in the inducible fluorescence production after the plateau at 1000 nM CdII. Therefore, when testing Hg or Cd bioavailability with respect to an environmental variable, we suggest using 5 nM for HgII and 300 nM for CdII.
In some instances, signal production can be properly attributed to Hg or Cd bioavailability, but in other cases, signal production can be affected by variation in the physiological state of the biosensor cell host (e.g., the metal of interest or environmental conditions tested are toxic). In Figure 4A, 5 nM Hg bioavailability was tested over a gradient of Zn (0-10 µM). In both Hg-inducible and constitutive strains, there is a similar decrease in signal with increasing Zn concentrations. Therefore, one cannot discriminate whether the signal results from lowered bioavailability or is a result of Zn toxicity. In Figure 4B and 4C, 5nM Hg bioavailability was tested over a gradient of MgII (0-10 mM) and MnII (0-10 µM). Increasing MgII and MnII concentrations decreased the fluorescence signal of the inducible strain. On the other hand, the constitutive strain did not show a decrease in fluorescence with increasing MgII and MnII concentrations (MgII and MnII are beneficial for the cells in the production of the FbFp, as demonstrated by an increase in the fluorescence signal). This demonstrates that the cells are viable and the fluorescence decrease of the Hg-inducible strain results from a decrease in Hg bioavailability. This data emphasizes how important it is for all biosensor assays to also provide constitutive measurements of overall cell fitness.
Figure 3: Linear ranges of the biosensor with Mercury and Cadmium. Maximum fluorescence measured as relative fluorescence units (RFU) ± 1 Standard Deviation emitted by E. coli NEB5α harboring the pUC57merR-Pp (Hg-Inducible) and pUC19Balch-Pp (Constitutive) with the addition of A) HgII (0-15 nM) and B) CdII (0-1,000 nM) under anaerobic conditions. Fluorescence was the average of 3 technical replicates at 37 ˚C. Please click here to view a larger version of this figure.
Figure 4: Example of an inconclusive result with Zinc and a conclusive result with Magnesium and Manganese. Maximum fluorescence measured as relative fluorescence units (RFU) ± 1 Standard Deviation emitted by E. coli NEB5α harboring the pUC57merR-Pp (Hg-Inducible) and pUC19Balch-Pp (Constitutive) with the addition of A) ZnII (0-10 µM), B) MgII (0-10 mM), and C) MnII (0-10 µM) under anaerobic conditions. [HgII] was set to 5 nM for all treatments and fluorescence was the average of 3 technical replicates at 37 ˚C. Please click here to view a larger version of this figure.
Microbial biosensors are useful tools to identify mechanisms by which metals are interacting with microbes. Here, we describe a method that can anaerobically quantify HgII and CdII bioavailability to a Gram-negative host cell (E. coli) and give a quantifiable result within a few hours. One of the major strengths of this protocol is that it allows extensive control of metal speciation in the exposure medium by avoiding strong binding ligands or components that may lead to metal precipitation. Metal speciation has been modelled and tested in this exposure medium using PHREEQC17, however other metal speciation software may be deployed. In the case of no added ligands, Hg speciation is expected to be present as 97% Hg(OH)2 and 3% Hg(NH3)22+, while Cd speciation will be present as 59% Cd-β-Glycerophosphate, 25% Cd2+, and 16% CdSO4. Using a simple thermodynamic modelling software, the user can design exposure media and test for the bioavailability of the metal of interest. In addition, the biosensor host cell (E. coli NEB5α) is viable over a wide range of pH (5-8.5) and NaCl concentration (0-0.55 M)17.
Hg methylation is an anaerobic process, and the protocol outlined in this study does not have a requirement for oxygen, allowing for more accurate description of anaerobic metabolism on metal bioavailability. This is important because the presence of oxygen alters gene expression profiles48,49 and hence, potential transport pathways; therefore this method presents an advantage over currently exising aerobic alternatives. The biosensing construct presented here can potentially be used with other anaerobic hosts that may be more relevant for mercury methylation (e.g., Geobacter, Desulfovibrio), but maybe less tractable than E. coli. One current limitation of the approach presented here is that our limit of detection has not yet reached pM levels, contrary to existing aerobic systems4,19,37. It is however important to note that to achieve these low detection limits several steps need to be taken44: i) ligand addition is required to ensure that Hg remains in solution and does not adsorb onto the microbial cell wall (Hg will be irreversible bound to cell surface thiols preventing its bioavailability25,27; see the threshold response for Hg in Figure 3A), ii) modifications to cell density, or iii) modify the genetic construct to include transport proteins of the mer-operon (namely merT and merP), increasing Hg flux inside the cell50,51. These modifications would be beneficial in detecting low concentrations of Hg, but not necessarily ideal when assessing environmentally relevant situations. Whereas previous cadmium biosensors primarily exist as a "proof of principle", they were designed in complex media that do not allow the investigator to assess the role of speciation on bioavailability41,42,43,44.
The biosensor is an incredibly useful tool in determining mechanisms in which metal species are bioavailable. Because the host organism is not a Hg-methylator, it may only be used to develop a model for how Hg may enter Gram-negative bacteria and not a definitive rationale for how Hg-methylators acquire Hg. Other methods exist for determining Hg bioavailability, such as methylation, as an outcome of uptake or the use of a mass balance approach10,15,20,45,46. That being said, the method presented here offers the advantage of quasi real time bioavailability data in viable cells. We do not recommend that this method be used to quantify total Hg or Cd levels in an environmental matrix. Despite the proposed use of biosensors to determine metal concentrations in the environment36, many more readily available standard methods are available such as ICP-MS, FAAS (for Cd analysis) or Cold vapor atomic absorption spectroscopy (for Hg analysis). The biosensor can however be used to determine if a given environmental matrix has the potential to enhance or hamper bioavailability; this is achieved by performing standard additions.
The pH of the exposure media may be altered to anywhere within the range of 5 and 8.5, provided the MOPS Free acid (buffer) is exchanged with an alternate free acid of a buffer with the appropriate pKa (please see list of appropriate buffers (Ferreira et al. (2015)47) and adjusted with KOH to the appropriate pH when making the exposure media. In addition, the method is not limited to Hg and Cd, but could be extended to other metals using other transcription regulators.
The exposure assay may be modified to explore the influence of other electron acceptors such as O2 and fumarate on Hg or Cd bioavailability. Minor modifications to the method to utilize both O2 and fumarate as electron acceptors are available upon request.
In summary we would like to emphasize the following points: I) It is imperative that the concentrations of Cd or Hg stocks are known in step 2, as these will be used to calibrate the biosensor. II) On the exposure day, the growth of cells must be stopped at an OD600 of 0.6 (± 0.1) and that care is taken when resuspending the cell cultures, as the biosensor is calibrated to this cell density. III) Lastly, it is important that the exposure medium is made meticulously on the exposure day. To ensure the success of the protocol, multiple cultures should be grown simultaneously (to circumvent the possibility of growth failure) and the growth medium should be remade weekly (to circumvent the metastability of the media and possible contamination). It should also be noted that biological replicates (multiple cell cultures) express variability when it comes to signal production. Although the fluorescent responses may vary from culture to culture, the fluorescent trends in response to a given variable should remain the same throughout numerous biological replicates.
The authors have nothing to disclose.
We would like to thank comments from two anonymous reviewers as well members of the Poulain Lab for insightful discussion on the development of the anaerobic biosensor. An Early Researcher Award from the Province of Ontario and a Discovery Grant and an accelerator supplement from the Natural Sciences and Engineering Research Council of Canada to A.J.P. funded this study.
|7 ml standard vial, rounded interior
|Vial tray, 21 mm openings
|4 for (4 x 8 grid)
|24 mm Closure
|LID CORNER NOTCH BLK STR CS/50
|Corning 96- well clear-bottom nonbinding surface microplate
|Anaerobic Chamber (glovebox)
|The air in the anaerobic chamber should be (97 % N2 ± 2 % and 3 % H2 ± 2 %)
|Converts O2 to H2O in the anaerobic chamber. Not required but recommended.
|450 (±10) nM filter for the microplate reader
|500 (±10) nM filter for the microplate reader
|Anaerobic culture vial (balch tubes) + rubber stoppers
|Modified for anaerobic culture tubes
|MA-3000 (mercury analyzer)
|Any pH probe will work
|50 mL conical sterile polypropylene centrifuge tubes.
|0.22 µm polyethersulfone syringe filter + syringe
|Sterile/clean glass bottles
|For growth media and standards
|Sterile/clean plastic or PTFE bottles
|For alkali solutions (NaOH/KOH)
|Reagents powders: Na2MoO4, Na2SeO4, H3BO3, NaOH, KOH, MnSO4, ZnSO4, CoCl2, NiCl2, ampicillin sodium salt, Difco M9 Minimal Salts, Glucose, MgSO4, Thiamine HCl, NaNO3, l-leucine, l-isoleucine valine, EDTA sodium salt, FeSO4, Sodium Beta-Gylcerophosphate, Mops free acid, (NH4)2SO4, Hg(NO3)2, CdCl2
|Sterile Milli-Q water
|Autoclaved or filter sterilized is fine.
|Analytical grade H2SO4
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