3.1 The optimum time of cell incubation and Mn(II) exposure for Mn(III) accumulation by strain LLDRA6
The accumulation of unstable Mn(III)-intermediate is an essential, indispensable step for generation of Mn(IV) oxides from Mn(II) ions, also indicative of the occurrence of Mn(II) oxidation. To determine the effect of cell activity on Mn(III) accumulation, and when Mn(III) accumulation will occur in the presence of strain LLDRA6, trapping assays of Mn(III)-intermediate by the ligand PP were performed on cells with different incubation times and different Mn(II) exposure times.
Figure 1(A, B, C and D) presents the UV-vis adsorption spectra of bacterial suspensions consisted of Mn(II), PP and cells at incubation times of 4 h, 8 h, 12 h, and 24 h, respectively. As a frequently-used chelating agent, PP has shown to be capable of stabilizing the transient Mn(III)-intermediate, thus forming Mn(III)-PP complexes that are easily detected by UV-vis spectrophotometer (Cui et al. 2008). It can be seen that characteristic peaks of Mn(III)-PP complexes formed under different incubation times all occur at 258 nm, which are similar to that of Mn(III)-PP complexes generated by Pseudomonas putida MnB1 (Thi et al. 2018). As cell incubation time increased from 4 h to 24 h (Figure 1A-D), the absorbance intensities at 258 nm gradually declined, suggesting that before Mn(II) exposure, cells with 4 h of incubation might be the most active for production of Mn(III)-intermediate in comparison to cells with longer incubation time. An explanation for this trend could be that at 4 h of incubation, cells were in the mid-exponential growth phase, and then they entered in the stationary growth phase since the 12th h, as evidenced by OD600 in the previous study (Li et al. 2020).
Furthermore, regardless of cell incubation time, 8 h of Mn(II) exposure was the optimum time for cells to accumulate Mn(III), since the corresponding absorbance intensity was always the strongest one at 258 nm. As the Mn(II) exposure time proceeded (> 8 h), the absorbances of Mn(III)-PP complexes at 258 nm unceasingly decreased, suggesting that the accumulated Mn(III) in cell suspensions was possibly oxidized to Mn(IV) oxides. It is worthy to note that at 4 h of Mn(II) exposure, the absorbance of 258 nm was far below that of 8 h of exposure, and was much closer to that of cells without Mn(II) exposure, indicating that Mn(III) accumulation by cells may start at as early as the 4th h and reach the plateau at the 8th h after the cells were exposed to Mn(II). Besides, it should be noted that cells without Mn(II) exposure had a background peak at 258 nm, not really meaning the presence of Mn(III).
Hence, the optimum time for Mn(III) accumulation by LLDRA6 at 4 h of cell incubation and 8 h of Mn(II) exposure were determined as the time parameters for cell incubation and the long-term Mn(II) exposure (i.e. a=4 and b=8 in section 2.4) in subsequent transcriptome sequencing, respectively (Fig. S2).
3.2 Transcriptomic differences of strain LLDRA6 between the short-term and the long-term exposure of Mn(II)
Prior to analysis of DEGs, RNA-seq data were validated by qRT-PCR. As shown in Fig. S3, for each selected gene, the log2-transformed mean values obtained from qRT-PCR showed the similar trends of transcriptional upregulation or downregulation to the log2-transformed fold changes from transcriptome data, suggesting the reliability of data from RNA-seq.
A total of 547 DEGs were identified between cells with short-term exposure (0.5 h) of Mn(II) (Mn(II)-short group) and cells with long-term exposure (8 h) of Mn(II) (Mn(II)-long group) (Table S3), while 1,468 DEGs were identified between cells with short-term incubation (CK-short group) and cells with long-term incubation (CK-long group) in the absence of Mn(II). Venn diagram analyses further showed that out of 547 DEGs, 300 were identified as unique ones between Mn(II)-short and Mn(II)-long groups, indicating that they were not caused by cell growth but only due to the exposure of Mn(II) (Fig. 2A). The other 247 DEGs were shared between control groups and Mn groups, suggesting that they were not only attributed to Mn(II) exposure but also attributed to cell growth.
In those 300 unique DEGs, 132 were upregulated and the other 168 were downregulated as shown in Table S4. To better understand their potential roles in Mn(II) oxidation after cells were exposed with Mn(II), KEGG pathway enrichment analyses were performed. The top 20 of KEGG pathways with the minimum P-value were shown as an enrichment bubble diagram (Fig. 2B). Notably, metabolism pathways related to aromatic or heterocyclic rings, such as phenylalanine metabolism (also known as phenylacetic acid metabolism), histidine metabolism, nitrotoluence metabolism, tyrosine metabolism, and ascorbate metabolism, were markedly enriched with Mn(II) exposure. Particularly, among them, phenylacetic acid metabolism aroused us a special attention, because its operon, consisted of fourteen-member phenylacetic acid genes (paa), was found to be the most transcriptionally upregulated in 547 DEGs between Mn(II)-short and Mn(II)-long groups (Table 1 and Table S3).
Table 1
Transcriptional expression profiles of 14 paa genes involved in phenylacetic acid metabolism pathway.
Gene ID
|
Orthology
|
CK-short_mean
TPMa
|
CK-long_mean
TPM
|
Mn(II)-short_mean
TPM
|
Mn(II)-long_mean
TPM
|
log2(CK-long/CK-short)
|
log2(Mn(II)-long/Mn(II)-short)
|
ProLLDRA6GL000272
|
paaY
|
32.64
|
61.73
|
74.01
|
758.54
|
0.91
|
3.35
|
ProLLDRA6GL000273
|
paaX
|
23.87
|
44.59
|
77.72
|
1562.21
|
0.90
|
4.32
|
ProLLDRA6GL000285
|
paaZ
|
12.84
|
23.63
|
31.08
|
1191.03
|
0.87
|
5.26
|
ProLLDRA6GL000277
|
paaH
|
112.91
|
561.12
|
66.61
|
4415.75
|
2.31
|
6.06
|
ProLLDRA6GL000282
|
paaC
|
9.93
|
27.88
|
20.71
|
2183.88
|
1.48
|
6.71
|
ProLLDRA6GL000278
|
paaG
|
13.17
|
121.74
|
36.80
|
3933.31
|
3.20
|
6.73
|
ProLLDRA6GL000279
|
paaF
|
1.84
|
18.44
|
12.12
|
1456.28
|
3.32
|
6.91
|
ProLLDRA6GL000281
|
paaD
|
1.04
|
3.16
|
5.16
|
656.06
|
1.59
|
6.98
|
ProLLDRA6GL000283
|
paaB
|
1.55
|
5.76
|
2.78
|
469.19
|
1.89
|
7.40
|
ProLLDRA6GL000284
|
paaA
|
1.25
|
3.07
|
5.01
|
1076.80
|
1.29
|
7.74
|
ProLLDRA6GL000275
|
paaJ
|
8.35
|
24.33
|
23.64
|
5117.34
|
1.54
|
7.76
|
ProLLDRA6GL000274
|
paaK
|
7.43
|
21.36
|
29.78
|
7525.53
|
1.52
|
7.98
|
ProLLDRA6GL000276
|
paaI
|
1.65
|
2.71
|
6.26
|
1671.92
|
0.71
|
8.05
|
ProLLDRA6GL000280
|
paaE
|
1.21
|
4.95
|
11.06
|
3051.35
|
2.03
|
8.10
|
a: TPM is the abbreviation for transcripts per kilobase million. |
Additionally, according to characteristics of enzymes catalyzing for Mn(II) oxidation (including copper binding motifs and heme/Ca(II) binding domains) as reported in other bacterial species, we found only two putative MCOs-type proteins possibly responsible for Mn(II) oxidation in strain LLDRA6, i.e. CotA and laccase (Table S5), after cluster of orthologous groups of proteins annotation (COG) for the complete genome of LLDRA6 (Su et al. 2013; Schlosser and Höfer, 2002). However, as shown in Table S5, in comparison with the short-term Mn(II) exposure, the transcriptional expressions of CotA genes were decreased after the long-term Mn(II) exposure. For laccase, there was no marked increase for transcriptional expression in response to the long-term Mn(II) exposure. So, under the current condition (50 mM of Mn(II) exposure), it cannot be determined that how much role have these MCOs-type enzymes played in the formation of Mn oxides via direct Mn(II) oxidation.
3.3 Sharp activation of phenylacetic acid metabolism pathway may hint ROS variation in LLDRA6
Previous studies have shown that in aerobic bacteria, the complex of enzymes encoded by the paa cluster is responsible for degradation of phenylacetic acid (PAA) and its derivatives (Teufel et al. 2010; Teufel et al. 2012; Cook, 2019). Intriguingly, after exhaustingly searching for relevant literatures, we find that phenylacetate catabolism is also important for resisting killing by H2O2 in bacteria (Green et al. 2020). A ∆paa mutant of Acinetobacter baumannii, which lacks part of the phenylacetate degradation pathway, was found to be more susceptible to H2O2 killing than a wild-type strain. The authors suggest that phenylacetate degradation may only be required to withstand environmental H2O2 encountered at high concentrations (≥ 40 mM) (Green et al. 2020). Herein, we supposed that for strain LLDRA6, the drastically upregulated expression of the paa cluster might attach to the variation of ROS contents in cells, due to the stress of ambient Mn(II) ions. Therefore, we further qualitatively analyzed variation of ROS contents for strain LLDRA6 with and without the exposure of Mn(II).
3.4 ROS variation in LLDRA6 with and without Mn(II) exposure
2′,7′-Dichlorofluorescin diacetate (DCFDA), capable of freely penetrating the cell membrane into the cytoplasm, is commonly used as a fluorometric probe for qualitative detection of various ROS in cells (Fan and Li, 2014). Since LB media could produce a certain amount of ROS (as later evidenced by Fig. 4 and Fig. 5), they were removed by centrifugation before detection of ROS by DCFDA. As shown in Fig. 3, fluorescent characteristic peaks of ROS produced by cells occur at 526 nm in the range of 510-650 nm. No matter which the Mn(II) exposure time was (0.5 or 8 h), the fluorescence intensities of cells with exposure of 50 mM Mn(II) were stronger than that of cells with no exposure of Mn(II), at an equivalent level of cell density (OD600 of 0.9 for the short-term exposure group, and OD600 of 1.7 for the long-term exposure group), suggesting a significant variation of ROS contents in strain LLDRA6 with and without the exposure of Mn(II).
3.5 Evidence for Mn(II) oxidation by LLDRA6 via superoxide generation coupled to hydrogen peroxide consumption
Commonly, ROS are short-lived oxygen radicals generated in the reduction of oxygen to water via the electron transfer chains, including
, H2O2, 1O2,
, HO−, ROO−, and RO−, etc. (Hansel et al. 2019; Fan and Li, 2014). As described by the reaction 1 above, since
and H2O2 play a key role in indirect oxidation of Mn(II), their contents in cells were specifically tested with and without Mn(II) exposure.
As shown in Fig. 4A, LB medium was able to generate a large amount of
by itself, even producing 2-fold greater amount of
in the presence of 50 mM of Mn(II). So, LB medium was removed by centrifugation prior to detection of
produced by cells.
Cells from both CK-short and CK-long groups were able to produce
(Fig. 4A). Interestingly, although the cell density of CK-long (OD600=1.7) was nearly two times greater than that of CK-short (OD600=0.9), the
concentration of cells from CK-long (2.19 µM) was much lower than that of cells from CK-short (5.26 µM). The similar trend of an inverse relationship between cell density and
production, has also been observed in other bacteria (Diaz et al. 2013; Hansel et al. 2019). These studies show that
concentrations steadily increased during active growth but then declined upon entering stationary phase, indicating their potential involvement in cell signaling except for simple responses to oxidative stress within microbial or environmental systems (Hansel et al. 2019).
When the cells were exposed to 50 mM of Mn(II) for 0.5 h,
concentration of cells from Mn(II)-short (5.06 µM) slightly declined, as compared to that of cells from CK-short (5.26 µM), perhaps indicating a rapid involvement of
in Mn(II) oxidation. As the Mn(II) exposure time increased to 8 h,
concentration of cells from Mn(II)-long significantly increased in comparison to that of cells from CK-long (from 2.19 to 2.88 µM), presumably because strain LLDRA6 was in middle of oxidizing Mn(II) ions when a vast amount of
were needed to constantly produce, as evidenced by the fact that 8 h of Mn(II) exposure was the optimum time for cells to accumulate Mn(III) (Fig. 1).
However, it cannot be ignored that substances other than living cells (e.g. LB media, Mn(II) ions and dead cells), were also capable of producing
during incubation (Fig. 4A). In order to confirm that
only generated by living cells is the true oxidant for Mn(II) oxidation, Mn(III)-trapping and leucoberbelin blue (LBB) stain assays (Krumbein and Altmann, 1973) were tested for living cells, LB media, and dead cells.
As shown in Fig. 4B, no absorption peak of Mn(III)-PP complexes could be found at 258 nm for LB medium in the absence and presence of Mn(II), showing that
produced by the medium was ineffective in oxidizing Mn(II). Also, dead cells in LB medium with or without addition of Mn(II) were not capable of producing Mn(III)-PP complexes after 8 h of incubation, since no characteristic peaks were detected at 258 nm. By comparison, living cells grown in LB medium with addition of Mn(II) produced a much stronger characteristic peak at 258 nm than living cells grown in the absence of Mn(II) did (considered as the background peak). Then, the strong characteristic peak produced by living cells grown in the presence of Mn(II), markedly attenuated as excessive superoxide dismutase (SOD) was used to scavenge superoxide. These results suggest that
only produced by living cells are involved in the formation of Mn(III)-PP complexes after 8 h of cell incubation. More importantly, after 4 d of continuous incubation, only the suspension of living cells incubated in LB medium with addition of Mn(II) showed deep blue in LBB stain assays (Fig. 4C), confirming the presence of Mn oxides. Thus, these results demonstrate that
only generated by living cells is the true oxidant for Mn(II) oxidation.
It is generally accepted that the production of
alone does not ensure the formation of Mn oxides in indirect Mn(II) oxidation, since H2O2 produced by cells may facilitate the reduction of Mn(III) to Mn(II) (Andeer et al. 2015; Lingappa et al. 2019). For instance, a marine alpha-protobacterium, Ruegeria sp. TM1040, does not generate Mn oxides despite its fast production rates of extracellular
(Diaz et al. 2013; Learman and Hansel, 2014).
As shown in Fig. 5, the concentration of H2O2 generated by cells after 0.5 h exposure of Mn(II) (Mn(II)-short) was much higher than that produced by cells from the control group (CK-short), perhaps owing to the rapid response to oxidative stress. As the exposure time increased to 8 h, the H2O2 concentration of Mn(II)-long cells significantly lowered in comparison to that of cells with no exposure of Mn(II) (CK-long), confirming a heavy consumption of H2O2 (from 25.99 to 22.87 µM), which facilitated to pull the equilibrium of the reaction 1 towards accumulation of Mn(III). A possible explanation for the consumption of H2O2 in cells of Mn(II)-long group, could be that the gene encoding catalase was significantly upregulated in transcription by 1.08-fold after the long-term Mn(II) exposure, as shown in Table S6, thus resulting in acceleration for decomposition reaction of H2O2.
Overall, these results suggest that Providencia sp. LLDRA6 exploited the indirect Mn(II) oxidation strategy of
production coupled to H2O2 consumption to form Mn oxides.
3.6 Prediction of links between ROS and phenylacetic acid metabolism pathway
Figure 6 presents a schematic diagram for the proposed process of Mn oxides generation by Providencia sp. LLDRA6. Regardless of the absence and presence of direct enzymatic oxidation in LLDRA6 (Table S5), the formation of extracellular Mn oxides by LLDRA6, as evidenced by our previous work (Li et al. 2020), now can be partially attributed to the attack of
that externally attached to cell membranes to Mn(II) (Learman et al. 2011b; Learman and Hansel, 2014; Andeer et al. 2015).
Once strain LLDRA6 is exposed to 50 mM of Mn(II), excessive Mn(II) will be harmful to the cell viability, as previously evidenced by OD600 (Li et al. 2020), probably through facilitating a higher level generation of ROS due to heavy-metal induced oxidative stress (Fig. 3) (Todsapol et al. 2020; Niu et al. 2020). ROS are often produced as by-products during the reduction of oxygen to water in electron transport chains (Hansel et al. 2019). Thus, enzyme complexes for respiration anchored in cell membrane may produce a large amount of extracellular
for reacting with Mn(II) to generate extracellular Mn oxides. In addition, some proteins, such as AHPs, malate dehydrogenase and xanthine dehydrogenase, are reported to take part in production of extracellular
via transmembrane transport (Andeer et al. 2015).
After the long-term exposure of Mn(II), the transcriptional level of the paa cluster is drastically upregulated (Table 1), indicating that PAA or PAA-like derivatives sharply accumulates and needs to be degraded as soon as possible, because from a healthy perspective, the excessive aromatic compounds are harmful to cell vitality (Teufel et al. 2010). So, the question arises, why does the paa cluster be transcriptionally activated in the long-term presence of Mn(II), or what the link might be between ROS and the activation of PAA catabolic pathway. According to the detailed information from catabolic pathway of PAA in aerobic bacteria as reported by Teufel et al. (2010), we find that the crucial process for degrading the inert aromatic ring of PAA, is the introduction of an oxygen atom into the aromatic ring catalyzed by the multicomponent enzyme PaaABCDE, and then the subsequent formation of a highly reactive oxygen-containing, seven-member heterocycle (named oxepin) catalyzed by PaaG (Fig. 6). Further, the ring of the C-O seven-member heterocycle is cleaved and opened at the oxygen site catalyzed by PaaZ.
Therefore, for inert, recalcitrant aromatic rings, the formation of highly reactive oxepins via epoxidation is an elaborate strategy for ring cleavage (Teufel et al. 2010). According to this conclusion, possible links between ROS and activation of phenylacetic acid catabolism could possibly be explained by three reasons as follows. (1) Commonly, the oxygen (O2) is used for epoxidation of the aromatic ring of PAA catalyzed by PaaABCDE. During the long-term exposure of Mn(II), the excessive ROS constantly produced in cells might replace the oxygen to involve in epoxidation, thus speeding up the catabolic pathway of PAA. (2) In fact, PAA catabolic pathway is not restricted to phenylacetate (Teufel et al. 2010). Other PAA-like aromatic compounds (e.g., benzoate, phenylalanine and styrene) can also enter in the same pathway for their degradation (Teufel et al. 2010). Hence, these PAA-like aromatic compounds might also be attacked by ROS to form highly reactive oxepins, thus greatly increasing their concentrations. Though exhibiting high chemical activity, oxepins are poisonous to cells, which need to be immediately hydrolyzed (Teufel et al. 2010), thereby further activating the PAA catabolic pathway. (3) the excessive oxygen radicals constantly produced in cells might result in a variety of oxygen-containing heterocycles that are similar to oxepins (oxepin-like ones) by attacking non-aromatic compounds (e.g., aliphatic hydrocarbons) (Meng et al. 2018), which may in turn greatly upregulate the transcriptional expression of the paa cluster.
Taken together, PAA, PAA-like derivatives and oxepin-like oxygen-containing heterocycles, may all be crowded in this catabolic pathway (Teufel et al. 2012), as a result of the response to Mn(II)-induced oxidative stress, consequently accelerating the transcription of the paa cluster. As to why does the bacterium choose PAA catabolic pathway for detoxification, a possible explanation is that all intermediates in this pathway are combined with CoA, which can be recognized rapidly and bound through CoA-binding motives of the processing enzymes (Teufel et al. 2010). More importantly, the final products of PAA catabolic pathway, acetyl-CoA and succinyl-CoA, are able to enter in the tricarboxylic acid cycle (TCA) for safe energy conservation.