Oxalate is the anion of a strong dicarboxylic acid, the oxalic acid (H2C2O4) that arises in the body from a combination of dietary sources principally of plant origin and endogenous synthesis.1
Oxalate (C2O42−) is the salt-forming ion of oxalic acid and can form oxalate salts combining with various cations, such as sodium, potassium, magnesium, and calcium. Approximately 75% of all kidney stones are composed primarily of calcium oxalate (CaOx)2 and hyperoxaluria, a condition involving high urinary oxalate concentration, is considered a primary risk factor for kidney stone formation, known as nephrolithiasis.
Nephrolithiasis is a common health disorder in the Western world, with a marked increase in prevalence over the past 20 years.
The explanation for the increasing incidence of nephrolithiasis in recent decades is still to be elucidated. Changes in lifestyle and dietary factors, such as increased consumption of animal proteins and salt, have been implicated as risk factors. Increasing prevalence of co-morbid conditions associated with kidney stones, including obesity and metabolic syndrome, may also have a role. Finally, recent reports suggest that climate changes associated with global warming may contribute to increasing rates of kidney stone formation.3–6
Nephrolithiasis causes significant morbidity among affected patients and a substantial financial burden because of lost work days and health care costs and is also associated with increased rates of chronic kidney disease and subsequent renal failure,7,8 hypertension, and myocardial infarction.9–11
Although it has been reported that dietary oxalate accounts for no more than 10% to 20% of oxalate excreted in the urine under normal conditions, more recent works12,13 have suggested that, even in the absence of gastrointestinal disorders, intestinal absorption of dietary oxalate can represent a more considerable contribution to urinary oxalate output.
Current therapeutic strategies often fail in their compliance or effectiveness, and CaOx stone recurrence is still common. Indeed, after an initial stone, there is a 50% chance of a second stone forming within 7 years if the condition is left untreated.14
Adequate fluid intake and dietary modifications are successful in the prevention of stone recurrence.15 Moreover, thiazide, potassium alkali, allopurinol, and tiopronin have been shown to be effective in the treatment of different stones. However, few novel therapies have emerged over the past decade and the continued poor compliance with drug therapy has led to a growing interest in new therapies aimed at preventing recurrent stone formation.
In particular, oxalate biotransformation by oxalate-degrading bacteria within the gastrointestinal tract is another key factor that could affect oxalate absorption, thereby decreasing the level of oxaluria.16–20
The best-known oxalate-degrading species is Oxalobacter formigenes, an anaerobic bacterium that inhabits the colon and depends solely on oxalate as a source of metabolic energy.21
Intestinal colonisation by O. formigenes begins in infancy, and by the age of 6 to 8 years, almost all children test positive for this bacterium.22 However, probably due to widespread antibiotic treatments, only 60% to 80% of adults test positive for O. formigenes. Moreover, a recent study demonstrated that black South Africans have a significantly higher faecal Lactobacillus spp. diversity and relative abundance of Bifidobacterium spp. compared with the higher risk of stone-forming typical of the white South African population.23
Because of their documented beneficial properties, Bifidobacterium and Lactobacillus strains are commonly used in dairy and pharmaceutical probiotic preparations and are generally recognised as safe for human consumption.24,25
The most widely accepted definition of probiotics states that they are “live microorganisms that, when administered in adequate amounts, confer a health benefit on the host.”26
The potential therapeutic application of lactobacilli and bifidobacteria in reducing hyperoxaluria in vivo through intestinal oxalate degrading activity is compelling and first reports are promising.27–31
In this study, we have screened different Lactobacillus and Bifidobacterium strains to investigate the capacity of some probiotic bacteria to degrade oxalate in vitro using reverse-phase high-performance liquid chromatography (HPLC).
Strains, Medium, and Growth Conditions
All bacterial strains used are shown in Table 1. All strains were stored at −80°C in MRS broth (Difco Laboratories, Milan, Italy) supplemented with 25% glycerol as cryoprotectant. Before experimental use, Lactobacillus strains were subcultured twice in MRS broth at 37°C under aerobic conditions for 24 hours. Bifidobacterium strains were subcultured twice in MRS broth supplemented with 0.5 g/L L-cysteine-hydrochloride at 37°C under anaerobic conditions for 24 hours.
O. formigenes (DSM 4420) has been employed as a positive reference to validate HPLC oxalate-degrading capability assay. O. formigenes was cultured according to the protocol described by Hungate et al32 in medium 419 (DSMZ, Germany) at 37°C for 48 hours, with the only exception that Na-oxalate was substituted with NH4-oxalate (Sigma-Aldrich, Milan, Italy).
Culture Conditions for Oxalate-degrading Tests
The oxalate degrading activity of 13 Lactobacillus and 5 Bifidobacterium strains was tested by HPLC analysis after growth in 10 mM ammonium oxalate medium.
Fresh bacteria were inoculated in a semi-defined medium (SM) as described by Campieri et al.33 The SM medium was composed of 5 mL of filtered sterilised ammonium oxalate solution (20 mmol/L ammonium oxalate and 40 g/L dextrose) added to 5 mL of base medium (protease peptone 20 g/L, yeast extract 10 g/L, Tween 80 2 mL/L, KH2PO4 4 g/L, Na-acetate 10 g/L, diammonium hydrogen citrate 4 g/L, MgSO4•7H2O 0.1 g/L, and MnSO4 0.1 g/L) previously sterilised at 121°C for 15 minutes. All reagents were supplied by Sigma-Aldrich (Milan, Italy) or Difco Laboratories (Milan, Italy).
Lactobacillus and Bifidobacterium strains and O. formigenes were added to SM to obtain an initial absorbance at 600 nm equal to approximately 0.100 OD (optical density) and then incubated anaerobically at 37°C for 24 hours. SM without the inoculum was used as control samples.
At the end of incubation period, samples were centrifuged at 6000g for 10 minutes to discharge the bacterial biomass. Supernatants were sterilised by filtration with Minisart 0.45 μm filters (Sartorius, Germany) and stored at −18°C.
Before HPLC analysis, supernatants were purified using Strata-X-A 33u Polymeric Strong Anion (500 mg/6 mL) SPE column (Phenomenex, UK) according to the manufacturer’s instruction. All reagents were supplied by Sigma-Aldrich (Milan, Italy).
The pH value of the supernatants was adjusted to 6 using 0.1 M NH3. The SPE column activation step was performed with 1 mL of methanol and conditioning step with 2 mL of 10 mM sodium formate. Subsequently, 1 mL of sample supernatant was loaded into the column. The washing step was performed with 1 mL of sodium acetate followed by 1 mL of methanol. The sample elution step was performed with 1 mL of 1M HCl and 1 mL of 3M HCl.
Before injection, the pH of each sample was adjusted to 6 using 0.1M NH3. All chemicals were HPLC grade and were supplied by Sigma-Aldrich (Milan, Italy).
Chromatographic analysis was performed with a Shimadzu HPLC system consisting of a CTO-20A column oven, a SPD-M20A UV visible diode array detector, a DGU-20A5 degaser, and a LC-20AT liquid chromatography pump. Peak areas were calculated using LabSolution LC single/PDA software.
The HPLC column used was a Synergi Hydro-RP column, 4 μm 250 mm×4.6 mm id. (Phenomenex, UK). Column cleaning and storage were performed according to the manufacturer’s instructions. All reagents used were of HPLC grade and provided by Sigma-Aldrich (Milan, Italy).
Oxalic acid was separated and quantified using the method described previously by Khaskhali et al,34 modified as follows. The mobile phase was composed of a filtered (Minisart 0.20 μm, Sartorius, Germany) and de-gassed solution of 20 mM potassium phosphate, buffered at pH 3 with orthophosphoric acid.
Aqueous oxalic acid standards were prepared in the range 0.02 to 20 mM, starting from a fresh stock solution of 200 mM oxalic acid. These solutions were stable for 3 months at 48°C.
Before analysis, the column was purged by pumping the mobile phase at 4 mL/min for 5 minutes and equilibrated with the mobile phase at a flow of 0.7 mL/min. The total cycle time was 35 minutes with a flow of 0.7 mL/min and 20 μL injection from each sample. The detector wavelength was fixed at 210 nm.
All reagents used were HPLC grade and provided by Sigma-Aldrich (Milan, Italy).
The results for oxalate degradation are expressed as mean±SD for the 3 different experiments. The statistical correlation between the percentages of oxalate degradation and OD values at 600 nm was evaluated using the Pearson index. By definition, the index could range between −1 and +1. A negative value means inverse correlation, whereas positive values represent direct correlation. If the index is between 0 and 0.3 (or −0.3), correlation is weak. If the index ranges between 0.3 and 0.7 (or −0.3 and −0.7), correlation is moderate. If the index is higher (or lower) than 0.7 (or −0.7), correlation is strong.
Quantification of Oxalate Degrading Activity
Lactobacilli were more efficient than Bifidobacterium strains in terms of oxalate degrading activity. L. paracasei LPC09 (DSM 24243) provided the best result, as 68.5% of ammonium oxalate was converted at the end of incubation, whereas the following best converters belong to the L. gasseri and L. acidophilus species (Table 1).
It is also interesting to point out that most bifidobacteria tested were only poor converters. B. breve BR03 was able to metabolise only 28.2% of the ammonium oxalate initially added to the medium.
In contrast, O. formigenes, used as positive control, was able to almost completely consume oxalate.
The Pearson correlation index between percentages of oxalate transformation and optical density at 600 nm was 0.276, thus meaning a weak direct correlation between the 2 variables.
Humans lack the enzymes needed to metabolise oxalate, and this potentially toxic compound is, therefore, managed in 3 different ways. It may be absorbed into the urinary tract and excreted in urine. Alternatively, oxalate in the gut can combine with calcium or magnesium, forming insoluble CaOx and/or magnesium oxalate that are eliminated with faeces. Furthermore, it can be degraded by microorganisms present in the gastrointestinal tract.
In light of the role of gut endogenous microbiota in metabolising oxalate, it is possible to hypothesise that the administration of oxalate-degrading microorganisms could be an alternative and innovative approach to reduce the intestinal absorption of oxalate and the resulting urinary excretion.
O. formigenes was the first oxalate-degrading obligate anaerobe to be described in humans35 and served as the paradigm organism in which anaerobic oxalate metabolism has been studied. Oxalate degradation by O. formigenes is important for human health, helping to prevent hyperoxaluria and disorders such as the development of kidney stones. Oxalate-degrading activity cannot be detected in the gut flora of some individuals, possibly because Oxalobacter is susceptible to commonly used antimicrobials.36
O. formigenes has 3 enzymes involved in the catabolism of oxalic acid. The process begins with the uptake of extracellular oxalate by the membrane associated with oxalate-formate antiporter, OxlT, encoded by the oxlT gene.37–39 Formyl-CoA transferase (Frc), encoded by the frc gene, activates the intracellular oxalate to form oxalyl-CoA.40,41 Oxalyl-CoA is then decarboxylated by the oxalyl-CoA decarboxylase enzyme (Oxc), expressed from the oxc gene. Formate and carbon dioxide are the end products,42 and the oxalate-formate antiporter, OxlT, catalyses the export of intracellular formate out of the cells.37,38 The decarboxylation of oxalate generates a proton pump gradient, which generates 1 ATP molecule when it is coupled with oxalate-formate transport.43O. formigenes uses oxalate as an exclusive source of energy and may, therefore, be described as a “specialist oxalotroph.”
Several studies have highlighted the oxalate degrading activity of different Lactobacillus and Bifidobacterium strains. However, it should be noted that these studies could not be directly compared from a quantitative point of view as there are several variations in the protocols used. For example, different sources and concentrations of oxalate (sodium vs. ammonium; range, 5 to 20 mM), different incubation times (1 to 5 days), cell preadaptation to oxalate, and different analytical protocols (enzymatic kits, HPLC, or capillary electrophoresis).
However, the general trends of the overall findings are of interest. Campieri et al33 identified potential probiotic strains through the evaluation of oxalate degradation by pure cultures of L. acidophilus, L. plantarum, L. brevis, S. thermophilus, and B. infantis. Among lactobacilli, L. acidophilus showed the highest percentage of metabolic breakdown of 10 mM ammonium oxalate (11.8%) and L. brevis the lowest (0.9%). S. thermophilus and B. infantis degraded 2.3% and 5.3% of oxalate, respectively. Weese et al44 also reported considerable variation in oxalate metabolism by different probiotics in vitro. They reported a mean oxalate degradation equal to 17.7% for 37 lactic acid bacteria (LAB) but without further identifying the strains. Turroni et al45 screened 14 Bifidobacterium strains for oxalate degradation activity. Among all the tested strains, no oxalate consumption was observed for B. adolescentis, B. bifidum, B. breve, B. catenulatum, B. longum subsp. longum and B. longum subsp. suis, whereas all the B. animalis subsp. lactis strains tested proved to be equally active, with 100% oxalate degradation. Moreover, the oxc gene was detected by PCR only in the B. animalis subsp. lactis strains.
Azcarate-Peril et al46 previously reported the presence of frc and oxc genes in L. gasseri and B. lactis. Turroni et al47 screened the oxalate-degrading ability of 60 Lactobacillus strains belonging to 12 species using an enzymatic assay. Among the tested strains, L. acidophilus and L. gasseri showed the highest activity, whereas the presence of oxc and frc genes, investigated through gene-specific PCR, was shown in all L. acidophilus and L. gasseri isolates able to transform >50% of oxalate.
This study involved a preliminary screening of the oxalate degrading activity in 13 lactobacilli and 5 bifidobacteria.
Our results suggest considerable variability in the ability of probiotic microorganisms to degrade oxalate in vitro, therefore confirming the strain-specific feature of many potentially beneficial effects of probiotics. The best converters of oxalate belong to the L. paracasei, L. gasseri, and L. acidophilus species.
The relatively low conversion rate observed with most bifidobacteria can probably be attributed to intrinsic oxalate toxicity toward this genus or to the lack of preadaption to oxalate before incubation. This hypothesis seems to be supported by the average slightly lower OD reading for all Bifidobacterium species tested compared with lactobacilli.
This aspect is mostly consistent with previously reported data from other groups.
There is a weak direct correlation between the growth extent of each strain and the ability to degrade oxalate added to the medium. In any case, a specific toxicity test would be required to better investigate this relationship, as a lower OD may indicate some negative influence exerted by ammonium oxalate itself on the ability of individual probiotic bacteria to grow in the SM medium.
Future studies in humans will be needed to investigate the ability of the best probiotic degraders to integrate into the autochthonous gut microbiota and reduce oxaluria, thus potentially preventing or decreasing the incidence and severity of kidney stone formation.
Our in vitro data confirm and extend a potential new perspective in the management of hyperoxaluria by providing an innovative and alternative tool to diminish the intestinal absorption of oxalate and the resulting urinary excretion.
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