Team:Wageningen UR/Wetlab/SyntheticDenitrification


iGEM Wageningen 2021

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Synthetic denitrification in Pseudomonas putida

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Synthetic denitrification in Pseudomonas putida

As part the ammonia pillar, the aim of this project was to build a synthetic denitrification pathway in P. putida, converting nitrate (NO3-) and nitrite (NO2-) into N2. To achieve this, the four denitrification enzymes, Nap, Nir, Nor and Nos were expressed in P. putida. We showed that the first two enzymes were functional individually as well as combined. Although the last two enzymes were successfully cloned and transformed into P. putida, no activity was observed yet.

Introduction

With Cattlelyst, we aim to completely convert ammonia (NH3) to dinitrogen gas (N2). This process is divided in two different processes, namely nitrification and denitrification. During nitrification NH3 is converted into nitrate (NO3-) and nitrite (NO2-) [1]. Denitrification is the reduction of NO3- or nitrite NO2- to the gaseous nitrogen compounds nitric oxide (NO), nitrous oxide (N2O) and N2 [2], [3]. Complete denitrification from NO3- to N2 is accomplished within four enzymatic steps (Figure 1) [4]. First, NO3- is converted to NO2- by nitrate reductase (Nap or Nar) and subsequently to NO by nitrite reductase (Nir). Then, NO is converted to N2O by nitric oxide reductase (Nor). Finally, nitrous oxide reductase (Nos) converts N2O into N2 [5], [6].

  • The difference between Nap and Nar
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    There are three types of prokaryotic nitrate reductases: (i) assimilatory nitrate reductase (nas), (ii) membrane-bound nitrate reductase (Nar) and (iii) periplasmic nitrate reductase (Nap). Nas converts NO3- into NO2-, which can subsequently be converted into NH3 used for biomass formation [7]. Nar and Nap are dissimilatory and convert NO3- into NO2-, which can subsequently be converted in NO. Nap is expressed under aerobic conditions and is required for aerobic denitrification [5].

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Figure 1: Schematic overview of enzymes and intermediates of the denitrification pathway. Abbreviations: nitrate reductase (Nap), nitrite reductase (Nir), nitric oxide reductase (Nor), nitrous oxide reductase (Nos), nitrate (NO3-), nitrite (NO2-), nitric oxide (NO), nitrous oxide (N2O) and dinitrogen gas (N2).

Since Cattlelyst operates with gas flows containing air, denitrification should function aerobically. In the last few decades, bacteria capable of combined heterotrophic nitrification and aerobic denitrification (HNAD) have been identified and studied. These bacteria can convert NH3 into N2 aerobically. Examples are Pseudomonas stutzeri [8], Pseudomonas aeruginosa [9] and Paracoccus denitrificans [10].

To ensure the optimal conversion of NO3- to N2 in the synthetic denitrification pathway, we employed two approaches:

  1. Plug and Play:

    2. Complete cloning of the denitrification pathway into P. putida. More about this approach can be found on the limiting nitrous oxide production in Pseudomonas putida page .

  2. Mosaic:

    3. Expressing all four denitrification enzymes separately in P. putida and comparing the enzymes from multiple native denitrifiers. This approach will be discussed on this page.

We expressed the genes from the native aerobic denitrifiers P. stutzeri JM300 [11], P. aeruginosa PA01 [9] and P. denitrificans DSM413 [12] in P. putida. Additionally, the genes from denitrifier Cupriavidus necator H16 were used. This is not a known aerobic denitrifier, but it does possess Nap, which is specifically known for being active in aerobic conditions [5]. By using the genes from different organisms, we were able to test and select the best enzymes for our pathway.

Approach

We created plasmids containing operons of the four different enzymes with all the hypothesized essential genes. An overview of all the genes can be found in ‘Cloning approach’. Once engineered and transformed into P. putida EM42 ΔnasT, the activity of the different enzymes was compared, when applicable.

  • Why P. putida EM42?
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    We chose the non-denitrifying organism P. putida as a chassis as it belongs to the same genus as the HNAD bacteria P. stutzeri and P. aeruginosa. P. putida KT2440 is recognized as a model laboratory bacterium due to its ability to adapt to harsh environmental conditions, fast growth, simple nutritional requirements and its extensive genetic toolbox [13]. Approximately 300 genes of P. putida KT2440 were eliminated, resulting in the derivative P. putida EM42 containing superior growth properties, a net increase of ATP availability and a higher final biomass yield [14]. P. putida EM42 was used as the chassis for the synthetic denitrification pathway.

  • Why ΔnasT?
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    To measure the activity of the expressed Nap, the amount of converted NO3- and produced NO2- should be quantified. However, P. putida EM42 contains an assimilatory nitrate reductase, which can convert NO3- into NO2-. This makes it difficult to distinguish between NO3- used for denitrification and NO3- used for growth. Nitrate and nitrite assimilatory reductases are regulated by the transcriptional regulator NasT [15], [16]. The encoding gene was deleted in P. putida EM178, resulting in a nitrate-blind strain unable to grow on NO3- as sole nitrogen source [17]. The same deletion was introduced in P. putida EM42 to ensure the same phenotypic output. This was established by homologous recombination with the assistance of the integrative vector pGNW [18].

  • Cloning approach
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    Each gene (Table 1) was individually PCR amplified from the genome of the organism and cloned into pSB1C3 with Golden Gate Assembly . Subsequently, the SEVA3.1 platform was used to assemble the genes in a SEVAb plasmid backbone, thereby introducing ribosome binding sites (BBa_J34801) and adding a promoter [19]. The resulting plasmids were transformed into E. coli DH5α competent cells, isolated and checked by sequencing. This was followed by transforming the plasmid into P. putida EM42 ΔnasT via electroporation. An overview of the constructs made can be seen in Table 2.

    Additionally, the nap operon from P. denitrificans was integrated in the genome of P. putida EM42 ΔnasT. This was required for testing the combination of Nap, Nir, Nor and accessory plasmid as only three antibiotics could be used simultaneously with P. putida and thus only three plasmids could be maintained. The integration of the nap operon was established by homologous recombination with the assistance of the integrative vector pGNW [20].

    Table 1: Overview of presumed necessary genes for the denitrification enzymes
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    Table 2: Overview of designed plasmids.
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Testing of constructs

ΔnasT

P. putida EM42 ΔnasT was tested for its ability to grow on NO3- or NO2- by growing it in M9 medium containing 30 mM acetate as carbon source (C-source). Different combinations of nitrogen sources (N-sources) were used, namely 5 mM NaNO3 or 2 mM NaNO2 with and without 2 g/L (NH4)2SO4. After growing for 24 hours, the NO3- or NO2- concentrations in the supernatant were quantified using the Nitrate Assay or the Griess Assay.

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Nap

To quantify the activity of Nap, P. putida EM42 ΔnasT containing the construct was cultured on M9 medium with 30 mM acetate as C-source) and 2 g/L (NH4)2SO4 as N-source supplemented with 5 mM NaNO3 as substrate for Nap. After 24 hours, the supernatant was taken and the NO2- concentration was quantified by performing a Griess Assay.

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Nir

To estimate the activity of Nir, P. putida EM42 ΔnasT containing the construct was cultured on M9 medium with 30 mM acetate as C-source and 2 g/L (NH4)2SO4 as N-source supplemented with 2 mM NaNO2 for Nir. After 24 hours, the supernatant was taken and the NO2- concentration was quantified by performing a Griess Assay. NO2- is known to be toxic for bacteria; therefore, a toxicity analyses was performed for NO2- on P. putida EM42 ΔnasT. The NO2- concentration was chosen according to this analysis.

It is important to note that the Griess Assay can also detect NO [50]. However, NO is very reactive due to an unpaired electron, which means NO has a short half-life in biological systems so little of the produced NO will be detected by the Griess Assay [51], [52]. Therefore, we designed a NO-responsive biosensor.

  • NO-biosensor
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    No accurate detection method was available to us for measuring NO. Therefore, a NO-responsive biosensor was designed to be able to detect the compound. The biosensor design is based on a NO-responding regulator (NorR) from C. necator [53]. When NO binds to NorR, transcription of the norAB promotor in C. necator is activated [54]. This mechanism can be adopted for the biosensor by combining NorR with a constitutive promoter (J23118), and by activating the transcription of GFP by the norAB promoter (Figure 2). Therefore, GFP is produced when NO is present; for example, by the conversion of NO2- by Nir. The backbone used for the biosensor were of the pSEVAbx3 category.

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    Figure 2: Schematic overview of our NO-responsive biosensor.
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Nor

As NO is a gas and also very toxic [38]. it was opted to use the NO donor sodium nitroprusside (SNP) for the screening of Nor. This donor has been used in experiments concerning denitrification or denitrifying bacteria before [55], [56] A toxicity analysis was performed for SNP on P. putida EM42 ΔnasT, and a SNP concentration was chosen accordingly. P. putida EM42 ΔnasT containing the Nor plasmid but without accessory plasmid was cultured in sealed bottles with air and M9 with 30 mM acetate as C-source and 2 g/L (NH4)2SO4 as N-source, supplemented with 1 mM SNP. N2O production was measured with gas chromatography - mass spectrometry (GC-MS).

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Nos

P. putida EM42 ΔnasT containing the Nos construct and the accessory plasmid was cultured in M9 with 30 mM acetate as C-source and 2 g/L (NH4)2SO4 as N-source in sealed bottles containing 21% oxygen and 0.5% N2O. To test whether the Nos construct is active in vivo, the depletion of N2O was measured with GC-MS. According to our GC technician, produced N2 was too difficult to measure with GC-MS, as N2 is abundant in air and minor differences cannot be measured.

Results

Nitrate- and nitrite–blind P. putida EM42

To create a nitrate- and nitrite- blind P. putida EM42, we deleted its nasT gene. Subsequently we checked for its ability to grow on NO3- and NO2- as nitrogen source (N-source), with or without NH4+. After culturing for 24 hours, NO3- and NO2- were not consumed by P. putida EM42 ΔnasT in either of the two conditions (Figure 3 and 4). In contrast, the wildtype was still able to consume these , when used as sole N-sources. When NH4+ is present in the media, NO3- and NO2- were not consumed by either the WT or the ΔnasT, meaning this knockout was not needed for the experimental design. However, it was still used in the following experiments.

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Figure 3: NO3- consumption by P. putida EM42 WT and P. putida EM42 ΔnasT when grown on 5 mM NO3- with or without 2 g/L NH4+ . Three biological replicates were used for each condition.
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Figure 4: NO2- consumption by P. putida EM42 WT and P. putida EM42 ΔnasT when grown on 2 mM NO2- with or without 2 g/L NH4+. Three biological replicates were used for each condition.
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Nap

We successfully cloned the nap operons of P. stutzeri, C. necator and P.stutzeri. Unfortunately, the operon from P. aeruginosa could not be completed, as napB was removed by P. putida numerous times. P. putida EM42 ΔnasT containing the Nap constructs were cultured with NO3- and the amount of NO2- produced was quantified and corrected for the change in optical density (OD600) (Figure 5). Both the Nap originating from C. necator and P. denitrificans worked in P. putida, but P. stutzeri Nap showed little activity. As P. denitrificans Nap showed the highest activity, we continued experiments with this enzyme.

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Figure 5: NO2- production by P. putida EM42 ΔnasT containing plasmid with nap operon originating from C. necator, P. stutzeri or P. denitrificans. The NO2- production was corrected for the change in OD over the time span of 24 hours. As control, P. putida EM42 ΔnasT containing pSEVAb22 without cargo was used. Three biological replicates were used for the control and Nap P. stutzeri and five biological replicates for C. necator and P. denitrificans Nap.

To be able to combine Nap with Nir and Nor and their accessory proteins, we integrated the nap operon from P. denitrificans in the genome of P. putida EM42 ΔnasT at the PP5322 site [57]. We compared the activity of this Nap with Nap expressed from the plasmid and we showed that the genomic Nap produced more than ten times more NO2- (Figure 6).

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Figure 6: NO2- production by P. putida EM42 ΔnasT containing nap operon originating from P. denitrificans on a plasmid (Nap plasmid) and integrated in the genome (Nap genomic). The NO2- production was corrected for the change in OD over the time span of 24 hours. As control, P. putida EM42 ΔnasT containing pSEVAb22 without cargo was used. Three biological replicates were used for the control and five biological replicates for Nap.
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Nir

There are two different types of Nir enzymes: heme d1-containing (NirS) and copper-containing (NirK) [28] NirS enzymes were divided into two operons/plasmids in our approach: a catalytic operon and a heme d1-producing operon. Previous research has shown that all heme d1-producing genes are necessary for NirS to be active [31]. Neither of the catalytic and heme d1-producing operons of P. stutzeri were ligated or cloned successfully. For P. aeruginosa, only the catalytic operon was cloned successfully. Nevertheless, this strain was not tested for NO2- consumption as the heme d1-producing operon was not available. From the four native denitrifiers, only P. stutzeri possess a NirK enzyme. The operon for this Nir was successfully cloned into P. putida EM42 ΔnasT.

The Nir containing strains were cultured with NO2- and the amount of NO2- consumed was estimated. The change in optical density (OD600) was also determined. While the OD of the control group (P. putida EM42 ΔnasT) increased normally in 24 hours, the OD of the Nir strains decreased. However, we did see signs of growth: a clear pellet was found in the samples after centrifugation at the 24-hour point, while non was visible at the start of the experiment. Moreover, the medium was cloudier after 24 hours. Figure 7 shows the consumption of NO2- over the 24-hour timespan. A notable difference between the control and the Nir strains can be seen. Approximately 20% of the added NO2- is converted in 24 hours. Unfortunately, the NO-responsive biosensor could not be used to prove the production of NO. Nevertheless, these results are a clear indication Nir is active in P. putida.

  • NO-biosensor
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    While the NO-responsive biosensor should work in theory, we were not able to get it working physically. The main problems occurred in the cloning of the construct. The norR gene gained a point mutation in its sequence, which resulted in the introduction of a stop codon in the middle of the sequence. Within the timeframe of iGEM 2021, we were unable to remove the mutation and transform the construct in P. putida. For future research, we recommend finishing and testing the biosensor to confirm NO is produced by Nir.

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Figure 7: NO2- consumption by P. putida EM42 ΔnasT containing the plasmid with the copper dependent nir operon originating from P. stutzeri. The NO2- production was not corrected for the change in OD over the time because the OD of the Nir containing strain decreased over the span of 24 hours. As control, P. putida EM42 ΔnasT. Three biological replicates were used for the control and test strain.

Both enzymatically active Nap and Nir were successfully cloned in P. putida EM42 ΔnasT. Therefore, we decided to test them simultaneously to see whether the enzymes can work consecutively. The Nap and Nir plasmids were cloned into P. putida EM42 ΔnasT to create P. putida EM42 ΔnasT pNap + pNir. This strain was grown with NO3- (5 mM) for 24 hours. Figure 8 shows the results of the growth experiment. Clearly, more NO2- is produced with Nap alone in comparison to Nap combined with Nir. This indicates that the produced NO2- from Nap is consecutively consumed by Nir. Unfortunately, the NO-responsive biosensor could not be used to confirm the production of NO by Nir. Nevertheless, these results are a clear indication that the enzymes are simultaneously functional.

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Figure 8:NO2- production by P. putida EM42 ΔnasT containing the nap operon originating from P. denitrificans on plasmid (Nap) or containing the nir operon origination from P. stutzeri on plasmid as well (Nap + Nir). The NO2- production was corrected for the change in OD over the time span of 24 hours. As control, P. putida EM42 ΔnasT containing pSEVAb22 without cargo was used. Three biological replicates were used for the control and five biological replicates for Nap and Nap + Nir.
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Nor

For Nor, only the operon with P. stutzeri genes was successfully ligated and cloned. The Nor operon was ligated with two different ORI (RSF1010 and RK2) and three different promotors (J23100, J23105, and rhamnose inducible). Cloning both in E. coli DH5α competent cells or directly in P. putida EM42 ΔnasT gave problems: no positive colonies were detected, or big deletions of norC and/or norB, or in the promotor occurred. The only successful cloning of nor happened with the rhamnose inducible promotor in P. putida EM42 ΔnasT. The optimal rhamnose concentration (0.23 mM) was determined by performing a growth experiment in which the OD was measured continuously/repeatedly over time (Figure 9). Above 0.23 mM rhamnose, growth of the P. putida strain with Nor was impaired. All these difficulties in cloning and the rhamnose test suggest that the expression of Nor is toxic or straining for the bacteria. The NorC and NorB proteins are membrane-localized in native denitrifiers [5], which could be causing these issues in cloning.

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Figure 9:Induction of nor operon in P. putida EM42 ΔnasTwith different rhamnose concentrations. Three biological replicates were used for P. putida EM42 ΔnasT containing pSEVA22-NorCBD (Nor).

The activity of the successfully cloned Nor operon was tested in sealed bottles with SNP (1 mM) and rhamnose (0.23 mM). No production of N2O was detected. This Nor plasmid was also transformed into the P. putida EM42 ΔnasT:nap together with the Nir plasmid to create a strain first three denitrification enzymes (P. putida EM42 ΔnasT:nap + pNir + pNor). This strain was also tested in sealed bottles, but nitrate (2 mM) was added instead of SNP. Unfortunately, no N2O was detected in this experiment either. We also were able to ligate and clone the accessory operon nirQOP in a separate strain. The accessory operon was also transformed into the P. putida EM42 ΔnasT:nap + pNir + pNor strain to test whether the accessory genes were not strictly accessory genes and more so essential for successful Nor activity. The accessory operon was the cargo of a pSEVAb43 backbone. This backbone contains a streptomycin resistance gene. Sadly, P. putida is known for acquiring natural resistance to this antibiotic relatively easily [58]. When we transformed our P. putida EM42 ΔnasT:nap + pNir + pNor strain with the accessory plasmid, we sadly only gained resistant strains. Therefore, a new strategy to test Nor with its accessory genes is needed. Within the timeframe of the iGEM 2021 competition this was no longer possible. Nevertheless, testing Nor with accessory genes is still promising.

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Nos

Multiple attempts have already been made to heterologously express Nos in vivo [44], [45]. Although in these studies Nos originating from P. stutzeri was expressed in E. coli and P. putida and was active in vitro, the challenge remains to make it work in vivo. Appropriate electron donors are required to couple the electron transfer chain to Nos. Therefore, we created an accessory plasmid containing multiple putative electron donors for Nos (Table 1). Both the plasmids containing the nos operon from P. stutzeri and the plasmid containing the electron donors were successfully cloned and transformed into P. putida EM42 ΔnasT. A rhamnose inducible promoter was used for the nos construct, as the same cloning problems were encountered as with nor. We determined the optimal rhamnose concentration (0.23 mM) by performing a growth experiment in which the OD600 was measured continuously over time (Figure 10).

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Figure 10: Induction of nos operon in P. putida EM42 ΔnasTwith different rhamnose concentrations. P. putida EM42 ΔnasT containing pSEVA22-NosRZDFYLTatA and pSEVA83- ApbE-blue copper protein 1 – blue copper protein 2 – c550 (Nos). As control, P. putida EM42 ΔnasT containing pSEVAb22 without cargo was used (Control). Three biological replicates were used for the control and two biological replicates for Nos.

We tested the activity of Nos by culturing this strain with N2O and subsequently measuring the reduction in N2O with GC-MS. Unfortunately, no N2O consumption was observed. The same experiment was performed with P. stutzeri JM300 to see if this native aerobic denitrifier could convert the N2O into N2, but also no reduction of N2O was observed. This could indicate that the experimental conditions were not optimal for Nos activity, but within the time frame of iGEM it was not possible to repeat this experiment.

Conclusion

We successfully expressed the first two enzymes of the synthetic denitrification pathway, namely Nap and Nir, in P. putida EM42 ΔnasT. We showed that Nap was able to convert NO3- into NO2- and Nir was able to convert NO2- into NO. Moreover, when combining these enzymes in P. putida EM42 ΔnasT, no NO2- accumulation was observed. This suggests that the enzymes can work consecutively. Functional expression of the remaining two enzymes of the pathway, was not established. Therefore, more research is needed to determine whether the enzymes could be active in P. putida.

  • References
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    1. C. JD and L. KM, “Nitric oxide is an obligate bacterial nitrification intermediate produced by hydroxylamine oxidoreductase,” Proc. Natl. Acad. Sci. U. S. A., vol. 114, no. 31, pp. 8217–8222, Aug. 2017, doi: 10.1073/PNAS.1704504114.
    2. G. Giannopoulos et al., “Tuning the modular Paracoccus denitrificans respirome to adapt from aerobic respiration to anaerobic denitrification,” Environ. Microbiol., vol. 19, no. 12, pp. 4953–4964, 2017, doi: 10.1111/1462-2920.13974..
    3. N. Shearer, A. P. Hinsley, R. J. M. Van Spanning, and S. Spiro, “Anaerobic growth of Paracoccus denitrificans requires cobalamin: Characterization of cobK and cobJ genes,” J. Bacteriol., vol. 181, no. 22, pp. 6907–6913, 1999, doi: 10.1128/jb.181.22.6907-6913.1999..
    4. J. Guo et al., “Pathways and organisms involved in ammonia oxidation and nitrous oxide emission,” Crit. Rev. Environ. Sci. Technol., vol. 43, no. 21, pp. 2213–2296, 2013, doi: 10.1080/10643389.2012.672072..
    5. B. Ji et al., “Aerobic Denitrification: A Review of Important Advances of the Last 30 Years,” Biotechnol. Bioprocess Eng., vol. 20, pp. 643–651, 2015, doi: 10.1007/s12257-015-0009-0.
    7. J. Yang et al., “A critical review of aerobic denitrification: Insights into the intracellular electron transfer,” Sci. Total Environ., vol. 731, p. 139080, 2020, doi: 10.1016/j.scitotenv.2020.139080..
    8. L. JT and S. V, “Nitrate assimilation by bacteria,” Adv. Microb. Physiol., vol. 39, pp. 1–30, 1998, doi: 10.1016/S0065-2911(08)60014-4..
    9. M. Miyahara et al., “Potential of aerobic denitrification by pseudomonas stutzeri TR2 to reduce nitrous oxide emissions from wastewater treatment plants,” Appl. Environ. Microbiol., vol. 76, no. 14, pp. 4619–4625, Jul. 2010, doi: 10.1128/AEM.01983-09..
    10. Z. Zhu et al., “Quorum sensing systems regulate heterotrophic nitrification-aerobic denitrification by changing the activity of nitrogen-cycling enzymes,” Environ. Sci. Ecotechnology, vol. 2, p. 100026, Apr. 2020, doi: 10.1016/J.ESE.2020.100026..
    11. K. Medhi, A. Singhal, D. Chauhan, I. T.-B. technology, and undefined 2017, “Investigating the nitrification and denitrification kinetics under aerobic and anaerobic conditions by Paracoccus denitrificans ISTOD1,” Elsevier, Accessed: Aug. 07, 2021. [Online]. Available: https://www.sciencedirect.com/science/article/pii/S0960852417303565?casa_token=JEH58HEeh00AAAAA:lh6s3jAtkFGbj2_y1hjnfYRW-lXdM6UIDAEYWdJM9UO1YDDGjgNZjzakaFbkaj_k5K4SlHOjiQ.
    12. L. Wittorf, C. M. Jones, G. Bonilla-Rosso, and S. Hallin, “Expression of nirK and nirS genes in two strains of Pseudomonas stutzeri harbouring both types of NO-forming nitrite reductases,” Res. Microbiol., vol. 169, no. 6, pp. 343–347, Jul. 2018, doi: 10.1016/j.resmic.2018.04.010.
    13. Z. Qu, L. R. Bakken, L. Molstad, Å. Frostegård, and L. L. Bergaust, “Transcriptional and metabolic regulation of denitrification in Paracoccus denitrificans allows low but significant activity of nitrous oxide reductase under oxic conditions,” Environ. Microbiol., vol. 18, no. 9, pp. 2951–2963, 2016, doi: 10.1111/1462-2920.13128.
    14. M. Martin-Pascual et al., “A navigation guide of synthetic biology tools for Pseudomonas putida,” 2021, doi: 10.1016/j.biotechadv.2021.107732.
    15. E. Martínez-García, P. I. Nikel, T. Aparicio, and V. de Lorenzo, “Pseudomonas 2.0: genetic upgrading of P. putida KT2440 as an enhanced host for heterologous gene expression,” Microb. Cell Factories 2014 131, vol. 13, no. 1, pp. 1–15, Nov. 2014, doi: 10.1186/S12934-014-0159-3.
    16. A. Caballero, A. Esteve-Núñez, G. J. Zylstra, and J. L. Ramos, “Assimilation of nitrogen from nitrite and trinitrotoluene in Pseudomonas putida JLR11,” J. Bacteriol., vol. 187, no. 1, pp. 396–399, Jan. 2005, doi: 10.1128/JB.187.1.396-399.2005.
    17. V. M. Luque-Almagro, V. J. Lyall, S. J. Ferguson, M. D. Roldán, D. J. Richardson, and A. J. Gates, “Nitrogen Oxyanion-dependent Dissociation of a Two-component Complex That Regulates Bacterial Nitrate Assimilation *,” J. Biol. Chem., vol. 288, no. 41, pp. 29692–29702, Oct. 2013, doi: 10.1074/JBC.M113.459032.
    18. K. Hobmeier, H. Löwe, S. Liefeldt, A. Kremling, and K. Pflüger-Grau, “A Nitrate-Blind P. putida Strain Boosts PHA Production in a Synthetic Mixed Culture,” Front. Bioeng. Biotechnol., vol. 8, p. 486, May 2020, doi: 10.3389/fbioe.2020.00486.
    19. N. T. Wirth, E. Kozaeva, and P. I. Nikel, “Accelerated genome engineering of Pseudomonas putida by I-SceI―mediated recombination and CRISPR-Cas9 counterselection,” Microb. Biotechnol., vol. 13, no. 1, pp. 233–249, Jan. 2020, doi: 10.1111/1751-7915.13396.
    20. S. G. Damalas, C. Batianis, M. Martin-Pascual, V. De Lorenzo, and V. A. P. Martins Dos Santos, “SEVA 3.1: enabling interoperability of DNA assembly among the SEVA, BioBricks and Type IIS restriction enzyme standards,” doi: 10.1111/1751-7915.13609.
    21. N. T. Wirth, E. Kozaeva, and P. I. Nikel, “Accelerated genome engineering of Pseudomonas putida by I-SceI―mediated recombination and CRISPR-Cas9 counterselection,” Microb. Biotechnol., vol. 13, no. 1, pp. 233–249, Jan. 2020, doi: 10.1111/1751-7915.13396.
    22. C. Coelho and M. J. Romão, “Structural and mechanistic insights on nitrate reductases,” Protein Sci., vol. 24, no. 12, pp. 1901–1911, Dec. 2015, doi: 10.1002/PRO.2801.
    23. P. Arnoux et al., “Structural and redox plasticity in the heterodimeric periplasmic nitrate reductase,” Nat. Struct. Mol. Biol. 2003 1011, vol. 10, no. 11, pp. 928–934, Oct. 2003, doi: 10.1038/nsb994.
    24. P. LC and C. JA, “Essential roles for the products of the napABCD genes, but not napFGH, in periplasmic nitrate reduction by Escherichia coli K-12,” Biochem. J., vol. 344 Pt 1, no. Pt 1, pp. 69–76, Nov. 1999, doi: 10.1042/0264-6021:3440069.
    25. C. Sparacino-Watkins, J. F. Stolz, and P. Basu, “Nitrate and periplasmic nitrate reductases,” Chem. Soc. Rev., vol. 43, no. 2, pp. 676–706, 2014, doi: 10.1039/c3cs60249d.
    26. N. E. Van Alst, L. A. Sherrill, B. H. Iglewski, and C. G. Haidaris, “Compensatory periplasmic nitrate reductase activity supports anaerobic growth of Pseudomonas aeruginosa PAO1 in the absence of membrane nitrate reductase,” Can. J. Microbiol., vol. 55, no. 10, pp. 1133–1144, Oct. 2009, doi: 10.1139/W09-065.
    27. N. A, B. THC, O. TW, R. DJ, L. ER, and C. JA, “The NapF protein of the Escherichia coli periplasmic nitrate reductase system: demonstration of a cytoplasmic location and interaction with the catalytic subunit, NapA.,” Microbiology, vol. 152, no. Pt 11, pp. 3227–3237, Nov. 2006, doi: 10.1099/MIC.0.29157-0.
    28. T. Brondijk, D. Fiegen, … D. R.-M., and undefined 2002, “Roles of NapF, NapG and NapH, subunits of the Escherichia coli periplasmic nitrate reductase, in ubiquinol oxidation,” researchgate.net, 2002, doi: 10.1046/j.1365-2958.2002.02875.x.
    29. S. Bali, D. J. Palmer, S. Schroeder, S. J. Ferguson, and M. J. Warren, “Recent advances in the biosynthesis of modified tetrapyrroles: The discovery of an alternative pathway for the formation of heme and heme d1,” Cell. Mol. Life Sci., vol. 71, no. 15, pp. 2837–2863, 2014, doi: 10.1007/s00018-014-1563-x.
    30. N. Hasegawa, H. Arai, and Y. Igarashi, “Need for Cytochrome bc1 Complex for Dissimilatory Nitrite Reduction of Pseudomonas aeruginosa,” Biosci. Biotechnol. Biochem., vol. 67, no. 1, pp. 121–126, 2003, doi: 10.1271/bbb.67.121.
    31. A. Jüngst, S. Wakabayashi, H. Matsubara, and W. G. Zumft, “The nirSTBM region coding for cytochrome cd1-dependent nitrite respiration of Pseudomonas stutzeri consists of a cluster of mono-, di-, and tetraheme proteins,” FEBS Lett., vol. 279, no. 2, pp. 205–209, 1991, doi: 10.1016/0014-5793(91)80150-2.
    32. S. Kawasaki, H. Arai, T. Kodama, and Y. Igarashi, “Gene cluster for dissimilatory nitrite reductase (nir) from Pseudomonas aeruginosa: Sequencing and identification of a locus for heme d1 biosynthesis,” J. Bacteriol., vol. 179, no. 1, pp. 235–242, Jan. 1997, doi: 10.1128/jb.179.1.235-242.1997.
    33. T. Klünemann, S. Henke, and W. Blankenfeldt, “The crystal structure of the heme d1 biosynthesis-associated small c -type cytochrome NirC reveals mixed oligomeric states in crystallo,” Acta Crystallogr. Sect. D Struct. Biol., vol. 76, pp. 375–384, 2020, doi: 10.1107/S2059798320003101.
    34. T. Klünemann, M. Nimtz, L. Jänsch, G. Layer, and W. Blankenfeldt, “Crystal structure of NirF: insights into its role in heme d 1 biosynthesis,” FEBS J., vol. 288, no. 1, pp. 244–261, Jan. 2021, doi: 10.1111/febs.15323.
    35. G. Layer, “Heme biosynthesis in prokaryotes,” Biochim. Biophys. Acta - Mol. Cell Res., vol. 1868, no. 1, p. 118861, 2021, doi: 10.1016/j.bbamcr.2020.118861.
    36. D. Helen, H. Kim, B. Tytgat, and W. Anne, “Highly diverse nirK genes comprise two major clades that harbour ammonium- producing denitrifiers,” BMC Genomics, vol. 17, no. 155, pp. 1–13, 2016, doi: 10.1186/s12864-016-2465-0.
    37. L. Philippot, “Denitrifying genes in bacterial and Archeal genomes,” Biochim. Biophys. acta (BBA)-Gene Struct. Expr., vol. 1577, pp. 355–376, 2002.
    38. M. Kahle, J. ter Beek, J. P. Hosler, and P. Ädelroth, “The insertion of the non-heme Fe B cofactor into nitric oxide reductase from P . denitrificans depends on NorQ and NorD accessory proteins,” BBA - Bioenerg., vol. 1859, no. 10, pp. 1051–1058, 2018, doi: 10.1016/j.bbabio.2018.05.020.
    39. W. G. Zumft, “Cell Biology and Molecular Basis of Denitrification,” Microbiol. Mol. Biol. Rev., vol. 61, no. 4, pp. 533–616, 1997.
    40. H. Arai, T. Kodama, and Y. Igarashi, “The Role of the nirQOP Genes in Energy Conservation during Anaerobic Growth of Pseudomonas aeruginosa,” Biosci. Biotechnol. Biochem., vol. 62, no. 10, pp. 1995–1999, 1998, doi: 10.1271/bbb.62.1995.
    41. A. Wüst, L. Schneider, A. Pomowski, W. G. Zumft, P. M. H. Kroneck, and O. Einsle, “Nature’s way of handling a greenhouse gas: the copper-sulfur cluster of purple nitrous oxide reductase,” Biol. Chem., vol. 393, no. 10, pp. 1067–1077, Oct. 2012, doi: 10.1515/HSZ-2012-0177.
    42. W. P and Z. WG, “Functional domains of NosR, a novel transmembrane iron-sulfur flavoprotein necessary for nitrous oxide respiration,” J. Bacteriol., vol. 187, no. 6, pp. 1992–2001, Mar. 2005, doi: 10.1128/JB.187.6.1992-2001.2005.
    43. U. Honisch and W. G. Zumft, “Operon Structure and Regulation of the nos Gene Region of Pseudomonas stutzeri, Encoding an ABC-Type ATPase for Maturation of Nitrous Oxide Reductase,” J. Bacteriol., vol. 185, no. 6, p. 1895, Mar. 2003, doi: 10.1128/JB.185.6.1895-1902.2003.
    44. T. LM, M. MA, D. DM, and C. V, “Structural studies of Apo NosL, an accessory protein of the nitrous oxide reductase system: insights from structural homology with MerB, a mercury resistance protein.,” Biochemistry, vol. 45, no. 40, pp. 12240–12252, Oct. 2006, doi: 10.1021/bi061089+.
    45. P. Wunsch, M. Herb, H. Wieland, U. M. Schiek, and W. G. Zumft, “Requirements for CuA and Cu-S center assembly of nitrous oxide reductase deduced from complete periplasmic enzyme maturation in the nondenitrifier Pseudomonas putida,” J. Bacteriol., vol. 185, no. 3, pp. 887–896, 2003, doi: 10.1128/JB.185.3.887-896.2003.
    46. L. Zhang, A. Wüst, B. Prasser, C. Müller, and O. Einsle, “Functional assembly of nitrous oxide reductase provides insights into copper site maturation,” Proc. Natl. Acad. Sci. U. S. A., vol. 116, no. 26, pp. 12822–12827, 2019, doi: 10.1073/pnas.1903819116.
    47. Z. L, T. C, A. SLA, and E. O, “The flavinyl transferase ApbE of Pseudomonas stutzeri matures the NosR protein required for nitrous oxide reduction,” Biochim. Biophys. acta. Bioenerg., vol. 1858, no. 2, pp. 95–102, Feb. 2017, doi: 10.1016/J.BBABIO.2016.11.008.
    48. S. Dell’Acqua, I. Moura, J. J. G. Moura, and S. R. Pauleta, “The electron transfer complex between nitrous oxide reductase and its electron donors,” JBIC J. Biol. Inorg. Chem. 2011 168, vol. 16, no. 8, pp. 1241–1254, Jul. 2011, doi: 10.1007/S00775-011-0812-9.
    49. “Pseudomonas stutzeri DSM 10701, complete genome - Nucleotide - NCBI.” https://www.ncbi.nlm.nih.gov/nuccore/CP003725 (accessed Oct. 18, 2021).
    50. M. K and H. T, “How does nitrous oxide reductase interact with its electron donors?--A docking study,” Proteins, vol. 59, no. 4, pp. 708–722, Jun. 2005, doi: 10.1002/PROT.20437.
    51. N. S. Bryan and M. B. Grisham, “Methods to Detect Nitric Oxide and its Metabolites in Biological Samples,” Free Radic Biol Med., vol. 43, no. 5, pp. 645–657, 2007.
    52. F. Cutruzzolà and N. Frankenberg-Dinkel, “Origin and impact of nitric oxide in Pseudomonas aeruginosa biofilms,” J. Bacteriol., vol. 198, no. 1, pp. 55–65, 2016, doi: 10.1128/JB.00371-15.
    53. S. Spiro, “Regulators of bacterial responses to nitric oxide,” FEMS Microbiol. Rev., vol. 31, no. 2, pp. 193–211, 2007, doi: 10.1111/j.1574-6976.2006.00061.x.
    54. A. Pohlmann, R. Cramm, K. Schmelz, and È. Friedrich, “A novel NO-responding regulator controls the reduction of nitric oxide in Ralstonia eutropha,” Mol. Microbiol., vol. 38, no. 3, pp. 626–638, 2000.
    55. A. Büsch, A. Pohlmann, B. Friedrich, and R. Cramm, “A DNA region recognized by the nitric oxide-responsive transcriptional activator NorR is conserved in β and γ-proteobacteria,” J. Bacteriol., vol. 186, no. 23, pp. 7980–7987, 2004, doi: 10.1128/JB.186.23.7980-7987.2004.
    56. K. Vollack and W. G. Zumft, “Nitric Oxide Signaling and Transcriptional Control of Denitrification Genes in Pseudomonas stutzeri,” J. Bacteriol., vol. 183, no. 8, pp. 2516–2526, 2001, doi: 10.1128/JB.183.8.2516.
    57. A. V Kwiatkowski and J. P. Shapleigh, “Requirement of Nitric Oxide for Induction of Genes Whose Products Are Involved in Nitric Oxide Metabolism in Rhodobacter sphaeroides 2.4.3*,” J. Biol. Chem., vol. 271, no. 40, pp. 24382–24388, 1996.
    58. J. E. Chaves et al., “Evaluation of chromosomal insertion loci in the Pseudomonas putida KT2440 genome for predictable biosystems design,” Metab. Eng. Commun., vol. 11, p. e00139, Dec. 2020, doi: 10.1016/J.MEC.2020.E00139.
    59. L. S. Van Overbeek et al., “Prevalence of streptomycin-resistance genes in bacterial populations in European habitats,” FEMS Microbiol. Ecol., vol. 42, no. 2, pp. 277–288, 2002, doi: 10.1016/S0168-6496(02)00345-8.
About Cattlelyst

Cattlelyst is the name of the iGEM 2021 WUR team. Our name is a mix of 1) our loyal furry friends, cattle, and 2) catalyst, which is something that increases the rate of a reaction. We are developing “the something” that converts the detrimental gaseous emissions of cattle, hence our name Cattlelyst.

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