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. 2024 Mar 22;19(4):1011–1021. doi: 10.1021/acschembio.4c00095

Discovery of a Gut Bacterial Metabolic Pathway that Drives α-Synuclein Aggregation

Lizett Ortiz de Ora †, Julia M Balsamo †, Kylie S Uyeda †, Elizabeth N Bess †,‡,*
PMCID: PMC11040608  PMID: 38517270

Abstract

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Parkinson’s disease (PD) etiology is associated with aggregation and accumulation of α-synuclein (α-syn) proteins in midbrain dopaminergic neurons. Emerging evidence suggests that in certain subtypes of PD, α-syn aggregates originate in the gut and subsequently spread to the brain. However, mechanisms that instigate α-syn aggregation in the gut have remained elusive. In the brain, the aggregation of α-syn is induced by oxidized dopamine. Such a mechanism has not been explored in the context of the gastrointestinal tract, a niche harboring 46% of the body’s dopamine reservoirs. Here, we report that Enterobacteriaceae, a bacterial family prevalent in human gut microbiotas, induce α-syn aggregation. More specifically, our in vitro data indicate that respiration of nitrate by Escherichia coli K-12, which results in production of nitrite that mediates oxidation of Fe2+ to Fe3+, creates an oxidizing redox potential. These oxidizing conditions enabled the formation of dopamine-derived quinones and α-syn aggregates. Exposing nitrite, but not nitrate, to enteroendocrine STC-1 cells induced aggregation of α-syn that is natively expressed in these cells, which line the intestinal tract. Taken together, our findings indicate that bacterial nitrate reduction may be critical for initiating intestinal α-syn aggregation.

Introduction

Although Parkinson’s disease (PD) has long been thought to originate in the brain, accumulating evidence indicates that some PD subtypes originate in the gastrointestinal (GI) tract.1,2 PD is characterized by motor impairment that arises when α-synuclein (α-syn) protein aggregates accumulate in dopaminergic neurons of the substantia nigra, the brain site of motor control;3 however, α-syn expression is not limited to the brain. α-syn is also expressed within the mucosa of the intestinal wall by enteroendocrine cells (EECs)4 as well as by enteric neurons that innervate the GI tract.5 At least 8 years prior to onset of motor symptoms in people with idiopathic PD, α-syn aggregates accumulate in GI tissue.6 These protein aggregates may subsequently propagate, putatively in a prion-like fashion, from the intestine to the brain via the vagus nerve that connects these organs.7,8 While there is evidence that intestinal α-syn aggregates foreshadow neurodegeneration in the brain, the molecular-level mechanisms responsible for intestinal α-syn aggregation have remained poorly understood.

Several lines of evidence suggest a microbial component in the development of α-syn aggregates and progression of PD. The gut microbiota is distinct in people with PD as compared to nondiseased controls.9−13 This dysbiosis is often characterized by an enrichment of the facultative anaerobic Enterobacteriaceae bacterial family whose abundance in the gut positively correlates with the severity of motor dysfunction in people with PD.9,14−16 Although it remains controversial whether gut microbiota dysbiosis is a cause or a consequence of PD pathogenesis, studies using mouse models implicate the gut microbiome in the etiology of this disorder. In germ-free mice overexpressing α-syn, GI-tract colonization using fecal samples from people with PD exacerbated motor deficits and brain pathology as compared to colonization using fecal samples from nondiseased controls.17 Additionally, induction of intestinal inflammation that is commonly associated with blooms in Enterobacteriaceae (DSS-induced colitis)18,19 resulted in accumulation of α-syn in GI tracts followed by the pathogenic buildup of this protein in the brains of α-syn-overexpressing mice.20,21

To identify specific gut bacterial biochemical processes that induce α-syn aggregation in the GI tract, we sought clues about characterized mechanisms of α-syn aggregation in the brain. In brain dopaminergic neurons, iron and dopamine can form a toxic pair that leads to aggregation of neural α-syn (Figure 1). Aging-related accumulation of iron in dopaminergic neurons causes oxidative stress that results in labile cytosolic ferrous iron (Fe2+) being oxidized to ferric iron (Fe3+).22 Cytoplasmic dopamine that is abundant in dopaminergic neurons can be readily oxidized by Fe3+ to highly reactive ortho-quinones.23 Dopamine-derived quinones interact with neural α-syn to cause misfolding that results in toxic α-syn oligomers.24

Figure 1.

Figure 1

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Upon oxidation of Fe2+ to Fe3+ in brain dopaminergic neurons, dopamine can be oxidized to ortho-quinones that cause α-syn to misfold and aggregate (figure created with BioRender.com, agreement number AA246MEBQ8.).

Like in the brain, the GI tract harbors dopamine and iron as well as expresses α-syn. Of the body’s dopamine pool, 46% is contained in the GI tract,25 and the gut microbiota is responsible for elevating the dopamine concentration in intestinal tissue.26 Iron, which can mediate dopamine oxidation, is present in high concentrations in the intestinal lumen (up to 25 mM), and increased dietary iron increases iron levels in intestinal cells.27 Although the oxidation state of labile cytosolic iron is predominantly Fe2+ in the nondiseased GI tract,28 conditions of oxidative stress increase the concentration of Fe3+, which could oxidize dopamine as depicted in Figure 1. With the convergence of concentrated dopamine and iron in the intestinal epithelium, as well as the expression of α-syn by intestinal EECs and enteric neurons, α-syn aggregation is poised to occur in the GI tract. We sought to identify gut bacterial biochemical processes that supply an oxidant capable of inducing iron-mediated dopamine oxidation and subsequent α-syn aggregation.

Although changes in redox potential that induce oxidative stress in the GI tract are typically associated with host metabolic processes,29−33 gut bacteria also modulate the redox potential of their environment.34 Here, we describe that the ability of Escherichia coli (a prototypic gut bacterium of the Enterobacteriaceae bacterial family18) to create an environment with an oxidizing redox potential is a stimulus that provokes iron and dopamine to cause α-syn aggregation. We identify that nitrite, which is an oxidant generated during Enterobacteriaceae nitrate dissimilatory metabolism that mediates oxidation of Fe2+ to Fe3+ in bacteria,18,35−37 stimulates a cascade of oxidation reactions that results in α-syn aggregation. Our results from in vitro experiments with both bacterial cultures and α-syn-expressing intestinal epithelial cells suggest a novel molecular mechanism by which the gut microbiota may influence PD pathogenesis.

Results

E. coli Nitrate Respiration Creates an Environment of Oxidizing Redox Potential

Given the positive correlation between PD severity and the abundance of Enterobacteriaceae in the gut microbiotas of people with PD,9 we sought to determine whether metabolic capabilities of this bacterial family are implicated in the pathogenic aggregation of α-syn. We were particularly intrigued by Enterobacteriaceae’s ability to perform anaerobic nitrate respiration, which results in production of an oxidant, nitrite.18,38,39 We hypothesized that an oxidizing redox potential would be created by nitrate-respiring Enterobacteriaceae and, thereby, stimulate shifts in the relative abundance of labile iron from being mainly Fe2+ (which putatively predominates in the reducing conditions of the gut microbiota40) to Fe3+ (which could subsequently oxidize dopamine and induce α-syn misfolding and aggregation).

To test our hypothesis, we anaerobically cultured E. coli K-12 to enable two types of metabolism: fermentation and respiration. Fermentation conditions were created by culturing E. coli K-12 in a minimal-nutrient medium supplemented with Fe2+ (500 μM) as well as glucose (20 mM) as the sole carbon source but without nitrate (media referred to as mM9–NO3). Conditions for nitrate respiration were generated by supplementing the same medium with nitrate (50 mM; media referred to as mM9+NO3). As shown in Figure 2a, supplementation with nitrate afforded a 1.6-fold increase in bacterial growth after 12 h of incubation as measured by optical density of cultures at 600 nm (OD600). These findings are consistent with reported in vivo findings: due to nitrate respiration being more energetically lucrative than fermentative metabolism,41 a higher concentration of intestinal nitrate enables a bloom in Enterobacteriaceae abundance in the gut microbiota.18

Figure 2.

Figure 2

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E. coli nitrate reduction generates an oxidant, nitrite, that creates an oxidizing redox potential in the bacterial environment and increases the relative abundance of Fe3+. E. coli K-12 was incubated in mineral media with nitrate (mM9+NO3) or without (mM9–NO3). (a) Growth was measured by optical density at 600 nm (OD600). (b) Oxidation–reduction potential (ORP) of bacterial cultures was measured, using an electrode, in culture media relative to sterile media. (c) Nitrite was quantified in cultures supplied with nitrate using the Griess assay. (d,e) Iron speciation was measured using the ferrozine assay in bacterial cultures where (d) media lacked nitrate (mM9–NO3) or (e) media was supplemented with nitrate (mM9+NO3). n = 3 biological replicates; bars denote means ± S.E.M.

In E. coli K-12 cultures, we also evaluated fluctuations in redox potential as a function of bacterial metabolism. A reducing redox potential (relative to sterile controls) was observed when bacteria was cultured in mM9–NO3 (Figure 2b). The progressively more reducing redox potential observed over the course of exponential bacterial growth is characteristic of fermentative metabolism that yields hydrogen, a reducing agent.42 In contrast, bacteria cultured in mM9+NO3 demonstrated a steadily increasing oxidizing redox potential (relative to sterile controls) over 18 h of incubation, reaching a maximum value of 308.17 ± 7.62 mV (Figure 2b). Notably, increases in the oxidative redox potential mirrored accumulation of nitrite in culture media (Figure 2c). Further supporting nitrite’s role as a redox-active metabolite, we observed that supplementation of sodium nitrite to sterile mM9 medium (mM9+NO2) afforded increases in redox potential in a concentration-dependent manner (Figure S1). In contrast, uninoculated sterile media supplemented with sodium nitrate (mM9+NO3) showed no significant variation in the redox potential of the solution. These findings support the notion that bacterial nitrate respiration can shift the redox potential of the environment to more oxidizing through the production of nitrite.

We next assessed whether the oxidizing redox potential afforded by nitrate-respiring conditions could shift the balance of iron speciation from favoring Fe2+ to favoring Fe3+. E. coli K-12 was cultured in mM9–NO3 or mM9+NO3 media, and the relative abundance of Fe2+ and Fe3+ was measured using the ferrozine assay.43 Under fermentation conditions that afforded a reducing redox potential (mM9–NO3), Fe2+ was the predominant iron species (Figure 2d). Conversely, under culture conditions for bacterial nitrate respiration (mM9+NO3), the oxidizing redox potential that corresponded to production of nitrite drove iron oxidation such that Fe3+ became the dominant oxidation state of iron (Figure 2e). Taken together, these findings demonstrate that the presence of nitrate enhances E. coli K-12 growth as well as creates an oxidizing redox potential that corresponds to nitrite production and results in the predominance of Fe3+ over Fe2+ in culture media.

E. coli Nitrate Respiration Initiates a Cascade of Oxidation Reactions that Lead to α-Syn Aggregation

Next, we sought to determine whether bacterial nitrate respiration could incite the cascade of oxidation reactions that are implicated in dopamine-dependent α-syn aggregation in cerebral dopaminergic neurons but that remain unexplored in the GI tract: Fe3+-mediated dopamine oxidation that forms ortho-quinones44−46 that cause α-syn to misfold and, subsequently, aggregate.23,24 To this end, we again anaerobically cultured E. coli K-12 until stationary phase (14 h) in either mM9+NO3 or mM9–NO3 but with the addition of α-syn monomer (20 μM) as well as dopamine (500 μM; mM9+NO3,+DA or mM9–NO3,+DA, respectively) or its vehicle (mM9+NO3,–DA or mM9–NO3,–DA, respectively). As before, bacteria cultured in nitrate-respiring conditions reduced the amount of nitrate to nitrite (Figure 3a). Accumulation of nitrite corresponded to an oxidizing redox potential (Figure 3b) and a shift in iron speciation so that the relative abundance of Fe3+ increased in comparison to cultures without nitrate supplementation (Figure 3c). Culture conditions in which nitrite was produced and dopamine was also present resulted in a lower relative abundance of Fe3+ as compared to conditions without dopamine. This finding is putatively a reflection of dopamine oxidation being coupled to the reduction of Fe3+, thereby increasing the relative abundance of Fe2+. Correspondingly, we were not able to measure redox potential in culture conditions that contained dopamine as redox potential did not stabilize under these conditions.

Figure 3.

Figure 3

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E. coli nitrate respiration instigates dopamine-dependent quinone formation and α-syn aggregation. E. coli K-12 wild-type or ΔmoaA was cultured for 14 h in mM9 media with α-syn monomer in the presence of nitrate (+NO3) or its absence (−NO3) and with dopamine (+DA) or without (−DA). Quantification of (a) nitrite (using Griess assay), (b) ORP relative to sterile media (using redox electrode), and (c) labile Fe3+ and Fe2+ (using ferrozine assay). (d–f) Representative images of (d) bacterial cultures as well as (e) membrane-stained quinone formation (using NBT stain) or (f) dot blots stained for α-syn aggregate formation (using immunostaining with MJFR-14 anti-fibril α-syn as the primary antibody). (g) Quantification of α-syn aggregates in dot blots. R.F.U.: relative fluorescence units. n = 3 biological replicates; bars denote means ± S.E.M.; significance was determined using ordinary one-way ANOVA with Sidak’s multiple comparisons test; and ****: P < 0.0001, ns: not significant.

In cultures containing dopamine, we observed formation of a dark pigment, which is characteristic of quinones (Figure 3d).47 Redox-cycling stain nitroblue tetrazolium (NBT), which specifically stains quinones,48 enabled detection of quinones in dot blots of nitrate-reducing bacterial cultures (Figures 3e and S2). Contrastingly, and as depicted in Figure 3d, no dark pigment was observed in dopamine-supplemented fermentative culture conditions (mM9–NO3,+DA) wherein Fe3+ relative abundance was significantly less than that in respiration culture conditions (mM9+NO3,+DA); correspondingly, no quinones were detected by NBT stain (Figure 3e; positive control shown in Figure S2). In the absence of dopamine supplementation to culture media, quinones were neither detected in nitrate-reducing nor in fermenting bacterial cultures. These data indicate that dopamine supplementation is necessary for quinone formation and that dopamine-dependent quinone formation occurs in the oxidizing conditions created upon bacterial reduction of nitrate.

Strikingly, dopamine-dependent quinone formation coincided with α-syn aggregation. Evaluation of α-syn aggregation was conducted by dot blot, immunostaining α-syn aggregates using antibody MJFR-14 on membranes spotted with culture media (Figure 3f; positive control is shown in Figure S3). Significantly, greater amounts of α-syn aggregates formed under nitrate-reducing conditions as compared to fermentative conditions—but only when dopamine was present (Figures 3g and S3). In the absence of dopamine, a culture environment of the oxidizing redox potential (mM9+NO3,–DA; Figure 3b) was not sufficient to induce α-syn aggregation (Figures 3g and S3). Nitrite-induced dopamine-dependent α-syn aggregation was recapitulated in abiotic experiments, in which nitrite was exogenously supplied rather than produced by bacteria (Figure S4). This finding was consistent regardless of the antibody used for detection: either MJFR-14 or 5G4, with the latter displaying enhanced selectivity for detecting α-syn aggregates relative to monomers (Figure S5).

We next used genetic knockout of nitrate respiration to further clarify the roles of nitrate and nitrite in initiating α-syn aggregation. First, we targeted a molybdenum cofactor (MoaA) that is incorporated into the active site of nitrate reductases and is essential for this enzyme to reduce nitrate to nitrite.49 MoaA was deleted from the wild-type E. coli K-12 to create the isogenic mutant E. coli K-12 ΔmoaA. Culturing E. coli K-12 ΔmoaA in mM9+NO3 media did not afford the growth advantage that was obtained when, upon culturing in the same media, the E. coli K-12 wild-type performed nitrate reduction (Figure S6). Moreover, E. coli K-12 ΔmoaA did not produce redox-active nitrite (Figure 3a), indicating that nitrate reduction was, indeed, inhibited by genetic deletion of moaA; likewise, a significantly less oxidizing redox potential was observed as compared to cultures of the wild-type strain cultured in mM9+NO3 media (Figure 3b). The less oxidizing redox potential of E. coli K-12 ΔmoaA cultures corresponded to significant reductions in the relative abundance of Fe3+ when E. coli K-12 ΔmoaA was cultured in either mM9+NO3,–DA or mM9+NO3,+DA as compared to analogous cultures of E. coli K-12 wild-type (Figure 3c). Without the ability of E. coli K-12 ΔmoaA to reduce nitrate and increase the oxidizing redox potential of the culture media, neither dopamine oxidation nor α-syn aggregation occurred (Figures 3e–g; S2 and S3). Taken together, these data indicate that the presence of nitrate alone does not induce α-syn aggregation; instead, we have demonstrated that bacteria that produce nitrite can transform an innocuous trio—Fe2+, dopamine, and α-syn monomers—into one that generates toxic α-syn aggregates.

Tungstate Inhibits α-Syn Aggregation Induced by Bacterial Nitrate Reduction

After identifying the instigating role of bacterial reduction of nitrate to nitrite in α-syn aggregation, we were curious about whether α-syn aggregation could be mitigated by chemically inhibiting bacterial nitrate respiration. To this end, we turned to tungstate, a chemical analogue of molybdate that renders Enterobacteriaceae nitrate reductases inactive.19 We evaluated the effect of sodium tungstate (0.5–100 mM) on E. coli K-12 (wild-type and ΔmoaA) cultured until stationary phase (14 h) in mM9+NO3 media supplemented with α-syn monomer (20 μM) and with or without dopamine (Figure S7). For E. coli K-12 wild-type cultures, a dose-response relationship was observed: increasing concentrations of sodium tungstate resulted in a decreasing production of nitrite (Figure 4a).

Figure 4.

Figure 4

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Tungstate limits α-syn aggregation by inhibiting bacterial nitrate reduction, which reduces redox potential of the bacterial environment. E. coli K-12 wild-type or ΔmoaA was cultured for 14 h in mM9 media with α-syn monomer in the presence of nitrate (+NO3), with dopamine (+DA) or without (−DA), and with sodium tungstate (WO42–) at varying concentrations (0–100 mM). Quantification of (a) nitrite (using Griess assay), (b) ORP relative to sterile media (using redox electrode), and (c,d) labile Fe3+ and Fe2+ (using ferrozine assay) in cultures (c) with dopamine or (d) without dopamine. (e–g) Representative images of (e) bacterial cultures of E. coli K-12 wild-type incubated in mM9+NO3,+DA supplemented with tungstate (0–100 mM) as well as (f) membranes stained for quinone formation (using NBT stain) or (g) dot blots stained for α-syn aggregate formation (using immunostaining with MJFR-14 anti-fibril α-syn as the primary antibody). (h) Quantification of α-syn aggregates in dot blots. R.F.U.: relative fluorescence units. n = 3 biological replicates; bars denote means ± S.E.M.; significance was determined using ordinary one-way ANOVA with Sidak’s multiple comparisons test; and ****: P < 0.0001, ***: P < 0.0007, **: P < 0.0026, *: P = 0.0168, ns: not significant.

In accordance with our findings that bacterial reduction of nitrate to nitrite creates a more oxidizing redox potential, progressive inhibition of this process using increasing concentrations of sodium tungstate supplemented to cultures of E. coli K-12 wild-type (in the absence of dopamine) correlated with decreasing oxidizing redox potentials of cultures (Figure 4b). As the redox potential of E. coli K-12 wild-type cultures decreased with increasing concentrations of tungstate, the relative abundance of Fe3+ also decreased in cultures with and without dopamine (Figure 4c) and without dopamine (Figure 4d). Tungstate (0.5–50 mM) inhibition of nitrate reduction by E. coli K-12 wild-type corresponded with decreased visible pigmentation of cultures (Figure 4e) and NBT-stained quinone (Figures 4f and S2) as well as decreased formation of α-syn aggregates (Figures 4g,h; S3). Although increasing tungstate concentrations from 0.5 to 100 mM resulted in significantly decreased nitrite level, relative redox potential, Fe3+ relative abundance, and α-syn aggregation, supplying E. coli K-12 wild-type cultures with up to 100 mM of tungstate did not ameliorate the effects of nitrite reduction to the extent that was observed upon genetic deletion of moaA. With 100 mM tungstate, nitrite concentration, Fe3+ relative abundance, and α-syn aggregates remained significantly greater in cultures of E. coli K-12 wild-type compared with E. coli K-12 ΔmoaA. Taken together, these results indicate that tungstate is a means to chemically limit (but not fully prevent) generation of the oxidizing environment created by E. coli nitrate reduction and, thereby, inhibit the cascade of oxidation reactions that leads to α-syn aggregation.

Redox-Active Nitrite Induces α-Syn Aggregation in Specialized Gut Epithelial Cells

We next examined the relevance of our proposed bacteria-induced α-syn aggregation mechanism to the mammalian gut. In the GI tract, α-syn is expressed by specialized epithelial cells called EECs;4 the dopamine metabolic pathway is also expressed by these cells.50 EECs are chemosensory cells at the interface between the gut luminal contents and the nervous system. While the apical side of these cells is in direct contact with the gut microbiome and its metabolites, a cellular projection (called a neuropod) on the basolateral surface of EECs forms synapses with enteric neurons, including those of the vagus nerve.51 Thus, EECs have been proposed as a potential site where environmental factors, including bacterial metabolites, could initiate α-syn misfolding and the prion-like cascade leading to PD.4,52

Owing to gut epithelial cells absorbing nitrite through passive diffusion,53 we hypothesized that this redox-active metabolite that is produced in the gut lumen by nitrate-respiring bacteria18,19 could induce aggregation of α-syn that is present in the cytoplasm of EECs.4 To test our hypothesis, we used murine STC-1 cells, an accepted model cell line for elucidating properties of native EECs.54 STC-1 cells were incubated with nitrate or nitrite (0.05–50 mM), and α-syn aggregation was analyzed via immunofluorescence staining using an antibody for α-syn fibrils (Figure 5a). STC-1 cells treated with 0.5 mM nitrate as compared with untreated cells showed no significant difference in amounts of α-syn aggregates (Figure 5b). In contrast, treatment with nitrite significantly induced α-syn aggregation in a concentration-dependent manner, with 0.5, 5, and 50 mM nitrite resulting in 2.1-, 12.9-, and 77.0-fold increases in aggregation, respectively, as compared to nitrate-treated cells (Figure 5b). Supplying cells with 0.05 mM nitrite afforded no significant amount of α-syn aggregation compared with nitrate-treated cells. Staining cells with an isotype control demonstrated no significant nonspecific background interactions (Figure S8). Taken together, these results not only provide strong evidence for our proposed model that nitrite induces α-syn aggregation within intestinal cells expressing this protein, but these findings also emphasize the importance of bacterial reduction of nitrate to nitrite to incite this pathogenic process.

Figure 5.

Figure 5

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Nitrite, but not nitrate, induces α-syn aggregation in enteroendocrine STC-1 cells. α-Syn aggregates per STC-1 cell upon incubation with nitrate (NO3–) or nitrite (NO2–) were (a) visualized (representative images) and (b) quantified using maximum intensity projections acquired by structured illumination microscopy (immunofluorescence staining of α-syn aggregates is shown in green; DAPI-stained cell nuclei are shown in blue). n = 3 independent biological replicates, 3–4 technical replicates for each; bars denote mean ± S.E.M.; significance determined by unpaired t-test; and *: P = 0.0113; ****: P < 0.0001, ns: not significant.

Discussion

Here, we show that the ability of Enterobacteriaceae, specifically E. coli, to modulate the redox potential of a bacterium’s environment plays a critical role in inducing the formation of α-syn aggregates. Microbial metabolism has been previously demonstrated to influence the redox potential of the environment;34 however, gut bacterial metabolic pathways are typically associated with creating more reducing environments, while the generation of oxidizing environments is commonly linked to host processes.29−33,55 Our data suggest that bacteria performing nitrate dissimilatory metabolism can generate an oxidizing environment. When this metabolic process occurs, Enterobacteriaceae reduce nitrate, a relatively redox-inert byproduct of the host inflammatory response,56 to nitrite, an oxidizing agent.

Using E. coli K-12 wild-type cultures, we demonstrated that the presence of nitrate in the growth medium results in its reduction to nitrite as well as the generation of a redox environment that is more oxidizing as compared to the same cultures without nitrate. In contrast, nitrate supplied to bacteria with a nitrate respiration defect (i.e., E. coli K-12 ΔmoaA) or to sterile media resulted in neither nitrite production nor a more oxidizing redox potential. These data indicate that E. coli K-12 nitrate metabolism generates an oxidizing environment. Notably, we showed that the shift in redox potential that accompanies nitrite production was sufficient to alter the relative abundance of iron species so that Fe3+ predominated over Fe2+ in cultures. This shift occurred in spite of anaerobic and reducing in vitro culture conditions similar to those of the GI tract that favor the prevalence of Fe2+ over Fe3+.40Enterobacteriaceae nitrate respiration may be an underappreciated mechanism by which gut bacteria disrupt their environment’s relative abundance of Fe2+ and Fe3+ as well as the metabolic processes mediated by this redox-active metal.

Iron has been implicated in PD onset due to the ability of Fe3+ to oxidize dopamine to ortho-quinones that cause α-syn monomers to aggregate.24,57,58 Through in vitro experiments, we demonstrated that gut bacterial nitrate reduction can induce the cascade of oxidation reactions that ultimately result in dopamine-dependent α-syn aggregation. This is the first report elucidating a gut bacterial metabolic pathway that directly influences α-syn aggregation in vitro. If conserved in the mammalian GI tract, this biochemical pathway may be a novel target for intervention strategies to prevent α-syn aggregation in the gut.

Due to tungstate’s inhibition of Enterobacteriaceae nitrate respiration,19 we sought to determine whether tungstate could be used to inhibit Enterobacteriaceae-induced α-syn aggregation. Tungstate exposure limited dopamine oxidation and α-syn aggregation in cultures of E. coli K-12 wild-type supplemented with nitrate. Additionally, tungstate treatment effectively lessened the oxidizing redox potential of the bacterial environment and increased the relative abundance of less-oxidizing Fe2+. Owing to the ability of oral tungstate treatment to effectively ameliorate murine colitis, which is exacerbated by gut bacterial nitrate respiration,19 the ability of tungstate to limit α-syn aggregation in vitro may have important therapeutic implications for limiting α-syn aggregation in the mammalian intestine.

Toward determining the significance of bacterial nitrate respiration to α-syn aggregation in the gut, we focused our efforts on EECs. EECs are emerging as a critical mediator of the gut–brain axis59,60 and have been implicated in PD as a source of intestinal α-syn.4,52 Since EECs can form synapses with the enteric nervous system,59 it has been hypothesized that α-syn aggregates may spread from EECs to the brain via the vagus nerve;4,52 however, the precise molecular stimuli of α-syn aggregation in EECs have remained elusive. Owing to our findings that nitrite induces dopamine-dependent α-syn aggregation in vitro, we suspected that nitrite could induce the same process in EECs. Within EECs, nitrite exposure (0.5–50 mM) afforded a dose-dependent increase in the amount of α-syn aggregation. Consistent with our in vitro experiments, the effect of nitrate (0.5 mM) on α-syn aggregation was no different than the sham treatment. Notably, the concentration of nitrate in the mucus layer of the intestinal lumen is on the order of 0.5 mM,18 which supports the physiological relevance of our findings. We demonstrated that nitrite—the product of gut bacterial Enterobacteriaceae nitrate respiration that occurs within the GI tract18,19—can induce α-syn aggregation in EECs, which line the GI lumen.

In summary, our data demonstrate that the gut microbiota, in particular E. coli (a prototypic organism of the Enterobacteriaceae bacterial family18), is capable of inducing α-syn aggregation. Enterobacteriaceae are more abundant in people with PD as compared to nondiseased, age-matched controls and are positively correlated with the severity of motor dysfunction;9 however, whether Enterobacteriaceae plays a causative role in PD has remained unknown. Here, we identified a specific metabolic pathway, bacterial nitrate reduction, by which Enterobacteriacaeae can generate the oxidative environment that causes dopamine oxidation and subsequent α-syn aggregation (working model depicted in Figure S9). While dopamine oxidation has been identified as a crucial component of α-syn aggregation mechanisms in the brain,24,57,58 we have demonstrated that dopamine-dependent mechanisms of α-syn aggregation are also likely relevant in the gut. Our findings that nitrite induces α-syn aggregation in EECs provide strong motivation for examining these cells as a likely conduit of the gut–brain axis in PD. Future work will focus on the ability of α-syn aggregates to spread to the enteric nervous system following their formation in EECs. If the cascade of reactions initiated by bacterial nitrate reduction and ending with α-syn aggregation is conserved in the mammalian gut, our findings position gut bacterial nitrate reduction as a novel target that may be leveraged for early intervention strategies to prevent intestinal α-syn aggregation and limit Parkinsonian neurodegeneration.

Materials and Methods

Bacteria and Culture Conditions

General

E. coli K-12 BW25113 was acquired from VWR (470179-082). E. coli ΔmoaA was constructed as previously described.61,62 Archived stocks of bacteria were maintained in 16% glycerol at -80 °C. Bacteria were routinely cultured in modified M9 mineral media (mM9, defined below) at 37 °C under anoxic and reducing conditions (2–5% H2, 20% CO2, with the balance being N2) in a COY anaerobic chamber. When indicated, mM9 media (described below) was supplemented with 50 mM sodium nitrate (Sigma, S5506-250G), 50 mM sodium nitrite (Fisher Scientific, M1065490100), 500 μM dopamine HCl (Alfa Aesar, A11136-06), and/or 20 μM α-syn (purified as described below). Prior to beginning experiments, the medium was equilibrated under anoxic conditions overnight to remove oxygen.

mM9 medium was prepared to contain the following: 1x M9 salts (Sigma, M6030-1KG), 2 mM magnesium sulfate (Fisher Scientific, 0338-500G), 100 μM calcium chloride (Fisher Scientific, AC219171000), 20 mM d-glucose anhydrous (VWR Life Science, Biotechnology grade), 0.2% (w/v) casamino acids (Fisher Scientific, DF0288-15-6), 1× vitamin supplement (ATCC, MD-VS), and 500 μM ferrous iron chloride (Oakwood, 098678-5g).

Purification of Recombinant α-Syn

Purification of α-syn was performed according to the method of Huang et al., 2005.63 Briefly, an overnight culture of E. coli Rosetta 2 transformed with pET-21a-α-syn (Addgene, 51485) was diluted 100-fold in LB supplemented with 100 μg/mL ampicillin. Expression of α-syn was induced for 5 h by adding 100 μM IPTG to the culture media when cell density reached an OD600 of 0.3–0.4. The cell pellet from a 1 L culture was resuspended in 100 mL of osmotic shock buffer (30 mM Tris–HCl, 40% sucrose, and 2 mM ethylenediaminetetraacetic acid disodium, pH 7.2) and incubated for 10 min at RT. The pellet collected by centrifugation at 12,000 rpm for 20 min was quickly resuspended in 90 mL of cold water followed by adding 37.5 μL of saturated MgCl2. The resuspension was kept on ice for 3 min. The supernatant containing periplasm proteins was collected by centrifugation at 3500 ×g for 20 min and dialyzed overnight against buffer A (20 mM Tris–HCl, pH 8.0). After centrifugation at 3500 ×g for 20 min, the supernatant was loaded onto a DEAE Sepharose column (GE healthcare) and eluted with a 0–0.5 M NaCl gradient in buffer A. The elution fractions were analyzed by SDS-15% PAGE, and the fractions containing only an 18 kDa band were combined for size-exclusion chromatography in 0.0095% MgCl2, 0.316% Tris–HCl, 0.58% NaCl, pH 7.5. Fractions containing pure α-syn were concentrated to 1 mg/mL, aliquoted, and stored at -20 °C.

Redox Potential Measurements

Redox potential was measured in 5 mL of bacterial cultures using a redox electrode with an Ag/AgCl reference electrode (Cole-Parmer, EW-59001-75). The values are reported relative to the standard hydrogen electrode (SHE) and were determined by measuring the offset of the reference electrode in a reference solution with a known redox potential (Thermo Scientific, 967961). Relative oxidation–reduction potential (ORP) was determined by subtracting the ORP value of the sterile control from the ORP of the experimental conditions.

Nitrate and Nitrite Measurements

To determine nitrate concentrations, the Griess-vanadium chloride method developed by Miranda et al. was used, with slight modifications.64 Briefly, 2 μL of bacterial culture was diluted in 198 μL of 1 M HCl. Of this suspension, 50 μL was mixed with 50 μL of a solution containing 0.1% (w/v) sulfanilamide (Sigma, S9251-100G) and 0.05% (w/v) N-(1-naphthyl)ethylenediamine dihydrochloride (Sigma, N9125-10G) in 0.5 M HCl followed by the rapid addition of 50 μL of 0.25% (w/v) vanadium(III) chloride (Sigma, 208272-1G) in 1 M HCl. The mixture was incubated at 37 °C for 30 min. Nitrite was evaluated in a similar manner except that samples were exposed to 50 μL of 1 M HCl instead of vanadium(III) chloride. In either case, following incubation, the absorbance at 540 nm was measured using a SPECTROstar Nano plate reader (BMG LABTECH). Sample concentrations were determined using calibration curves generated by measuring absorbance at 540 nm and performing linear regression against known concentrations of nitrate and nitrite prepared from sodium nitrate or sodium nitrite standards, respectively, at concentrations ranging from 1000 to 15.62 μM (two-fold dilution series).

Iron Speciation Assay

To determine the ratio of Fe2+/Fe3+ in media, we used the Fe2+-ferrozine assay as previously described, with slight modifications.43 Briefly, to determine the concentration of Fe2+ present in media, a bacterial culture (70 μL) was mixed with 70 μL of 50 mg/mL ferrozine (Hach, 230424) in 1 M potassium acetate buffer (pH 5.5). To determine the total iron present in the solution, ascorbic acid was used as a reducing agent to reduce Fe3+ to Fe2+. For this purpose, bacterial culture (90 μL) was mixed with 100 μL of 50 mg/mL ferrozine in 1 M potassium acetate buffer (pH 5.5) and 10 μL of 1 M ascorbic acid (Sigma, A92902-25G). Reduced and nonreduced samples were incubated at 37 °C for 30 min followed by centrifugation at 3000 ×g for 15 min. Then, 100 μL of the supernatant was used to determine the light absorption of the Fe2+-ferrozine complex at 562 nm using a SPECTROstar Nano plate reader (BMG LABTECH). Sample concentrations were determined using calibration curves generated by measuring absorbance at 562 nm and performing linear regression against the known concentrations of Fe2+ prepared from FeCl2 standards at concentrations ranging from 100 to 1.56 mM (two-fold dilution series). Fe3+ was calculated as the difference between total iron (the sum of Fe2+ and Fe3+ reduced to Fe2+) and Fe2+.

Nitroblue Tetrazolium Assay for the Detection of Quinones

The presence of quinones in bacterial cultures was evaluated as previously described.48 Briefly, under ambient conditions, 2 μL of each sample was spotted onto a polyvinylidene difluoride (PVDF) membrane (Fisher Scientific, ISEQ00005) that was previously activated in methanol and equilibrated in tris-buffered saline (pH 7.4). As a positive control, 2 μL of oxidized dopamine produced by Aspergillus tyrosinase (Worthington Biochemical, LS003789; detailed protocol described below) was used. The membrane was left to dry for at least 1 h. Then, the membrane was reactivated using methanol and submerged in a solution of 0.6 mg/mL NBT (Santa Cruz Biotechnology, sc-296003) in a potassium glycinate buffer (pH 10). The membrane was then incubated at RT, in the dark, for 45 min. After incubation, the membrane was washed twice in a solution of 0.16 M sodium borate (Sigma, S9640-25G) and photographed using an Epson V19 scanner.

Dopamine Oxidation Using Tyrosinase

Lyophilized Aspergillus tyrosinase (Worthington Biochemical, LS003789) was resuspended in 1000 μL of 50 mM phosphate buffer, 15% glycerol, pH 6.5, to create a 24 kU/mL solution (50 mg/mL, 390 μM). Single-use aliquots (100 μL) were prepared and stored at -20 °C. Aliquots were thawed on ice shortly before use. Dopamine (500 μM) was oxidized by adding 250 U (3.9 μM) of tyrosinase. The solution was incubated for 1 h at RT.

Immunodetection of α-Syn Aggregates

α-Syn aggregation was evaluated on methanol-activated PVDF membranes. Under anaerobic conditions, 2 μL of bacterial cultures were spotted and left to dry for at least 1 h. As controls, 2 μL of monomeric (Abcam, ab51189) and aggregated α-syn (Abcam, ab218817) were each spotted onto the membrane. The membrane was immunostained with anti-fibril α-syn (Abcam, ab209538; 1:50,000 dilution) as primary antibody and NIR 800CW donkey anti-rabbit IgG secondary antibody (LI-COR, 926-32213) according to the manufacturer’s instructions. The membranes were visualized using an Odyssey CLx imager (LI-COR) using the 800 nm channel. α-Syn aggregates were quantified using Image Studio data analysis software (LI-COR). Relative fluorescence unit (R.F.U.) values were scaled relative to sterile control samples, which were set equal to zero.

Abiotic Incubations

Samples with sodium nitrate (50 mM; Sigma), sodium nitrite (50 mM; Fisher Scientific), dopamine HCl (500 μM; Alfa Aesar), ferrous iron chloride (500 μM; Oakwood), and/or recombinant human α-syn monomer (20 μM; Abcam, ab51189) in double-deionized (DDI) water (36 μL) were prepared under anoxic and reducing conditions (2–5% H2, 20% CO2, with the balance being N2) in a COY anaerobic chamber. Samples were aliquoted into three 12 μL volumes. These triplicates were then incubated, in PCR tubes, at 37 °C for 4 h in the anaerobic chamber described.

Immunodetection of α-Syn Aggregates

α-Syn aggregation was evaluated in methanol-activated PVDF membranes. Under anaerobic conditions, 2 μL of each sample was spotted onto three different methanol-activated PVDF membranes. As controls, 2 μL (578 ng) of monomeric α-syn (Abcam, ab51189) and 578 ng of aggregated α-syn (Abcam, ab218817) were spotted onto each membrane. The membranes were allowed to dry for 1 h. One membrane was used as a quantitative control to ensure that the protein concentration in each sample was equivalent. The dried membrane was washed with DDI water before it was submerged in Ponceau stain, which had been prepared to contain Ponceau S (Sigma-Aldrich) and glacial acetic acid (Fisher Scientific). Ponceau stain was discarded after 1 min, and the membrane was subsequently washed with DDI water to remove excess water before visualizing. The other two membranes were blocked for 1 h using 5% Blotting Grade Blocker Non-Fat Dry Milk (Bio-Rad) in TBS. One of the blocked membranes was then probed with the MJFR-14 anti-aggregate α-syn primary antibody (Abcam, ab209538; 1:10,000 dilution) for 2 h before being incubated overnight at 4 °C. After four 5 min washes with TBS-T [0.1% Tween 20 (Sigma-Aldrich) in the prepared TBS], the membrane was subsequently immunostained with IRDye 800CW donkey anti-rabbit IgG secondary antibody (LI-COR, 926-32213; 1:20,000 dilution) for 45 min in the dark. Four more washes using TBS-T followed, before rinsing off the Tween 20 with TBS. The other blocked membrane was immunostained with the 5G4 anti-aggregate primary antibody (Millipore Sigma, MABN389; 1:4000 dilution) and IRDye 800CW donkey anti-mouse IgG secondary antibody (LI-COR, 926-32212; 1:20,000 dilution). The same steps as described above were followed for washing and immunostaining with the secondary antibody. The membranes were visualized by using an Odyssey CLx imager (LI-COR) on the 800 nm channel. Blots were quantified using Image Studio data analysis software (LI-COR) to analyze the relative aggregation of α-syn. Prism was used for further data processing.

Antibody Selectivity for α-Syn Aggregate versus Monomer

In DDI water, α-syn monomer (1 mg/mL; Abcam, ab51189) and α-syn aggregate (2 mg/mL; Abcam, ab218817) were each used to prepare separate dilution series containing the following amounts of protein: 350, 289, 144.5, 72.3, 36.1, 18.1, and 9.03 ng/μL. Of each sample, 2 μL was spotted in duplicate onto two different methanol-activated PVDF membranes, affording the following amounts of α-syn monomer or α-syn aggregate on the membranes in each dot: 700, 578, 289, 144, 72.3, 36.1, and 18.1 ng. Using the procedures described above, one membrane was immunostained using MJFR-14 anti-aggregate α-syn primary antibody (Abcam, ab209538; 1:10,000 dilution) followed by IRDye 800CW donkey anti-rabbit IgG secondary antibody (LI-COR, 926-32213; 1:20,000 dilution). The other membrane was immunostained using 5G4 anti-aggregate primary antibody (Millipore Sigma, MABN389; 1:4000 dilution) and IRDye 800CW donkey anti-mouse IgG secondary antibody (LI-COR, 926-32212; 1:20,000 dilution). The membranes were visualized using an Odyssey CLx imager (LI-COR) on the 800 nm channel. Blots were quantified using Image Studio data analysis software (LI-COR) to analyze relative aggregation of α-syn. Prism was used for further data processing.

Cell Lines and Growth Conditions

General

Enteroendocrine STC-1 (CRL-3254) cell line was obtained from the American Type Culture Collection (ATCC). Cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM, Corning) containing 4.5 g/L glucose, 2 mM l-glutamine, 10% (v/v) fetal bovine serum (Life Technologies), penicillin (100 U/mL), and streptomycin (100 μg/mL, Gibco). Cells were incubated at 37 °C in a humidified atmosphere with 5% CO2. Cells were serially passaged using 0.25% Trypsin–EDTA (Gibco). Cells were seeded onto two 8-well glass slides (Ibidi) at a density of 1 × 105 viable cells/well and incubated for 48 h. Then, the growth medium was replaced with fresh medium supplemented with either sodium nitrate, sodium nitrite, or vehicle. Cells were incubated for another 24 h before fixation.

Immunofluorescence

Fixation was performed using 10% formalin (Fisher Scientific, 22-170-402) for 20 min at RT. Cells were then permeabilized for another 20 min at RT using 0.1% Triton X-100 (Bio-Rad, 1610407) in phosphate-buffered saline (PBS), pH 7.4. After discarding the permeabilization solution, wells were washed with PBS, twice. Samples were then blocked for 1 h at room temperature in PBS containing 5% Normal Goat Serum (Thermo Scientific, 50197Z) and 0.2% bovine serum albumin (BSA; Fisher Scientific, 501613336) followed by washing three times using PBS. Subsequently, the glass slide chamber was incubated at 4 °C overnight with anti-α-syn aggregate primary antibody (Abcam, ab209538; 1:250 dilution), isotype control (Abcam, ab172730; 1:566 dilution), or vehicle. The chambers were then washed with PBS followed by an incubation with anti-goat Alexa Fluor-488 secondary antibody (Abcam, ab150077; 1:500 dilution) for 1 h at room temperature in the dark. After washing three times, ∼10 drops of VECTASHEILD PLUS Antifade Mounting Medium with DAPI (Vector Laboratories, H-1900) were added to each well. All demarcation and staining steps were performed at RT, but the chamber slides were stored at 4 °C in the dark.

Structured Illumination Microscopy

Images were acquired under identical conditions using a Zeiss Elyra 7 super-resolution microscope with a 63× oil immersion lens. Images were collected using 405 and 488 nm laser lines for excitation; emission filters used were BP 420-480 (DAPI) and BP 495-550 (AlexaFluor488). Sections were imaged in the absence of primary antibodies, and images were captured at the same gain as images with the primary antibody. No endogenous tissue fluorescence was observed in the absence of primary antibodies. Z-stack images were obtained and processed using SIM2 scaled to raw image. Quantification of the fluorescence signal for each sample was determined by obtaining the mean intensity of the maximum intensity projection for each image given by Zen Black 3.0 software. The number of cells in each image was counted, and mean intensity per cell was calculated. For each biological sample, approximately 30–40 cells were evaluated. The background fluorescence signal was accounted for each replicate by subtracting the mean intensity per cell for the respective untreated sample. Results are expressed as arbitrary units (a.u.) of mean intensity per cell.

Data and Statistical Analysis

Data were plotted and statistically analyzed using GraphPad Prism 8 software. Unless stated otherwise, means ± standard error of the mean (S.E.M.) were plotted. Statistical tests performed are specified in the figure legends. For all analyses, P ≤ 0.05 was considered significant. Sample sizes are noted within the main text and figure legends and within the Materials and Methods section.

Acknowledgments

The table of contents graphic was created with BioRender.com (agreement number DM240BYT0B). Financial support for this publication results from the University of California, Irvine School of Physical Sciences, the University of California Cancer Research Coordinating Committee (C21CR2124), and Scialog grants #28626 and #28648, sponsored jointly by Research Corporation for Science Advancement, the Frederick Gardner Cottrell Foundation, and the Paul G. Allen Frontiers Group. This study was made possible in part through access to the Optical Biology Core Facility of the Developmental Biology Center, a shared resource supported by the Cancer Center Support Grant (CA-62203) and Center for Complex Biological Systems Support Grant (GM-076516) at the University of California, Irvine.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acschembio.4c00095.

  • Additional experiment details, including redox potential measurements, full dot blot membranes, comparison of antibodies for detecting aggregated α-syn, growth data for bacterial cultures, isotype controls for immunofluorescence microscopy experiments, and a working model of bacteria-induced dopamine-dependent α-syn aggregation (PDF)

Author Present Address

The current affiliation of L.O.O. is the Department of Molecular Biology and Biochemistry, University of California, Irvine, California, USA 92617

Author Contributions

# L.O.O. and J.M.B. contributed equally. L.O.O. and E.N.B. developed the project. L.O.O., J.M.B., and E.N.B. designed experiments and performed data analysis. L.O.O. and J.M.B. performed experiments with contributions from K.S.U. L.O.O., J.M.B., K.S.U., and E.N.B provided critical feedback on experiments. L.O.O., J.M.B., and E.N.B. wrote the manuscript. E.N.B. acquired funds and provided project supervision and administration. All authors read and approved the final version of the manuscript.

The authors declare no competing financial interest.

Supplementary Material

cb4c00095_si_001.pdf (728.6KB, pdf)

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